Controlled tunnel gap device for sequencing polymers

ABSTRACT

The invention includes compositions, devices, and methods for analyzing a polymer and/or polymer unit. The polymer may be a homo- or hetero-polymer such as DNA, RNA, a polysaccharide, or a peptide. The device includes electrodes that form a tunnel gap through which the polymer can pass. The electrodes are functionalized with a reagent attached thereto, and the reagent is capable of forming a transient bond to a polymer unit. When the transient bond forms between the reagent and the unit, a detectable signal is generated and used to analyze the polymer.

REFERENCE TO RELATED APPLICATIONS

This application is a national stage application, filed under 35 U.S.C.§371, of International Application No. PCT/US2011/023185, filed Jan. 31,2011, which claims priority to U.S. Provisional Patent Application No.61/300,678, filed on Feb. 2, 2010, and to U.S. Provisional PatentApplication No. 61/378,838, filed on Aug. 31, 2010, both of which areincorporated herein by reference in their entirety.

GOVERNMENT RIGHTS

This invention was made with government support by grants HG004378 andR21HG004770, awarded by the National Institute of Health, by grantHG004378 from the Sequencing Technology Program of the National HumanGenome Research Institute, and by grant U54CA 143682 from the NationalCancer Institute. The government has certain rights in the invention.

BACKGROUND OF THE INVENTION

New approaches to DNA sequencing are required to reduce costs andincrease the availability of personalized genomics (M. Zwolak, M. DiVentra, Reviews of Modern Physics 80, 141 (2008)). In addition, longcontiguous reads would help to unravel the long-range structure of thegenome (E. Pennish, Science 318, 1842 (2007); A. J. Sharp, et al., Annu.Rev. Genomic Hum. Genet. ARI, 407 (2006). In contrast to Sangersequencing and next-generation methods, nanopore sequencing (D. Brantonet al., Nature Biotechnology 26, 1146 (2008)) is an enzyme-freetechnique in which DNA molecules are forced through a tiny apertureusing electrophoresis, so that a sequence-reading mechanism couldmaintain its fidelity over the entire length of a molecule. Ion currentthat passes through the pore is sensitive to the sequence in thenanopore (M. Akeson, et al., Biophys J. 77, 3227 (1999); A. Meller, etal., Proc. Natl. Acad. Sci. (USA) 97, 1079 (2000); N. Ashkenasy, et al.,Angew. Chem. Int. Ed. 44, 1401 (2005)) but all of the bases in thenanopore channel contribute to the current blockade (A. Meller, et al.,Phys. Rev. Lett. 86, 3435 (2001)) as well as those in the region of highfield beyond the pore (A. Aksimentiev, et al., Biophysical Journal 87,2086 (September 2004); M. Muthukumar, et al., Proc. Natl. Acad. Sci.(USA) 103, 5273 (2006)). In consequence, single base resolution has notyet been attained with an ion current readout. Lee and Thundat proposedthat electron tunneling across a DNA molecule might be localized enoughto sense and identify single nucleotides (J. W. Lee, and T. Thundat.U.S. Pat. No. 6,905,586 (2005)), a conjecture supported by thecalculations of Zwolak and Di Ventra (M. Zwolak, M. Di Ventra, NanoLett. 5, 421 (2005)). Further calculations show that thermal motion ofmolecules in the gap broadens the distribution of tunnel currents (J.Lagerqvist, et al., Biophys J. 93, 2384 (2007); R. Zikic et al., Phys.Rev. E 74, 011919 1 (2006)), reducing selectivity substantially. Therange of orientations of molecules in a tunnel gap can be greatlyreduced by using chemical bonds to tether it to the readout electrodes(X. D. Cui et al., Science 294, 571 (2001)), however, the use of strongbonds is not an option for DNA sequencing where the contact to theelectrodes must slide from one nucleotide to the next rapidly. Ohshiroand Umezawa demonstrated that hydrogen bonds can be used to providechemical contrast in scanning tunneling microscope images (T. Ohshiro,Y. Umezawa, Proc. Nat. Acad. Sci. 103, 10 (2006)) suggesting that theseweaker bonds can serve as “sliding contacts” to single molecules.

In applications WO2008124706A2 (“Sequencing by Recognition”), 61/037,647(Nanotube Nanopore for DNA Sequencing“), 61/083,001) (“Tandem Reader forDNA Sequencing.”) 61/083,993 (”Carbon Nanotube Based Device forSequencing Polymers“), 61/103,019 (”A Trans-base tunnel Reader forSequencing“), all of which are incorporated by reference, schemes forcontacting target bases in DNA in a tunnel gap with electrodesfunctionalized with reagents designed to hydrogen bond specifically toone base or another are described. In consequence, a different reader isrequired for each DNA base, so that a sequence has to be assembled byaligning the output of four separate readers. Furthermore, the relianceon reagents designed to target a specific site means that when twodifferent sites are targeted (one by each electrode) the electrodes haveto be functionalized independently, which is difficult to achieve in ananoscale gap.

SUMMARY OF THE INVENTION

The present invention provides compositions, devices, and methods foranalyzing a polymer and/or polymer unit. The polymer may be a homo- orhetero-polymer such as DNA, RNA, a polysaccharide, or a peptide. Thedevice has electrodes that form a tunnel gap through which the polymercan pass. The electrodes are functionalized with a reagent attachedthereto, and the reagent is capable of forming a transient bond to apolymer unit. When the transient bond forms between the reagent and theunit, a detectable signal is generated and used to analyze the polymer.The tunnel gap width is configured or adjusted to optimize selectivityof the signal generated when the electrodes form a transient bond to theunit of the polymer.

BRIEF DESCRIPTION OF THE FIGURES

FIG. 1 illustrates hydrogen bonding for T (FIG. 1A), G (FIG. 1B), C(FIG. 1C), and A (FIG. 1D) with electrodes functionalized with4-mercaptobenzamide.

FIG. 2 provides exemplary background tunneling signals in TCB for a biasof 0.5V. FIG. 2A is at a current of 10 pA and FIG. 2B is at a current of2 pA.

FIG. 3 shows the exemplary effect of electrode functionalization on thedistribution of current spikes for purines. Bare electrodes (FIG. 3A—dAand FIG. 3C—dG) give broad distributions (gap conductance 20 pS, 0.7 μMdA, 2.9 μM dG in TCB). Fits are Gaussian in the log of the current (seeFIGS. 18-20). Distributions narrow ten-fold when one electrode isfunctionalized with 4-mercaptobenzene (FIG. 3B—dA, FIG. 3D—dG) (gapconductance 12 pS, I_(bl)=6 pA, V=0.5V). Fits are to two Gaussians inthe log of the current with a peak at i₀ (“1”) and a second at 2 i₀(“2”) (see formula III). i₀=5.9 pA for dA and 5.6 pA for dG. When bothelectrodes are functionalized (FIGS. 3E—dA, 3G—dG) the peak currents areclearly different (i₀=9.4 pA for dG, i₀=16.5 pA for dA). FIG. 3F showsthe distribution for a mixture of dA and dG. The assignment of thehigher peak to dA is confirmed by the distribution measured with areduced concentration of dA (FIG. 3H). The high current tail in FIGS. 3Fand 3H is consistent with a small number of two molecule (dA+dG) reads.Distributions of the spike widths are given in FIG. 29.

FIG. 4 provides an exemplary plot of current vs. time trace for V=0.5V,background current=6 pA when adenosine diffuses into the gap. The insetshows a blow-up of a binding signal. Similar types of signals areobserved for all four nucleosides. See FIG. 15.

FIG. 5 shows the exemplary effect of electrode functionalization forpyrimidine reads. For reads with bare electrodes (broad distributions inFIGS. 5A and 5B) G_(bl) was increased to 40 pS to increase the countrate. The narrow distributions in 5A and 5B are taken with bothelectrodes functionalized and yield i₀=6.7 pA for dT and 13.3 pA for dC(G_(bl)=12 pS, I_(bl)=6 pA, V=0.5V). In a mixed solution, (FIG. 5C) thedT peak occurs at 8 pA and the dC peak occurs at 13.4 pA, an assignmentverified by measuring a mixture with half the concentration of dT (FIG.21).

FIG. 6 shows a exemplary distribution of current spikes for bothelectrodes when functionalized (dG, V=0.5 V, current=6 pA). A smallfraction of reads at twice the peak current for the main peak signalsimultaneous trapping of two molecules in the tunnel gap.

FIG. 7 provides a summary of exemplary reads. FIG. 7A shows peak currentfor dA (filled squares) and dT (filled circles) as a function of thebaseline conductance (at V=0.5V). Open squares (dA) and open circles(dT) show how the fraction of two molecule reads increases as the tunnelgap is made smaller. FIG. 7A shows that the measured molecularconductance increases linearly with G_(bl) (black circles dT, blacksquares dA, error bars are ±HWHH). The number of two molecule reads(open circles, dT, open squares, dA) increases at G_(bl)=20 pS, and theread rate is substantially reduced at G_(bl)=4 pS. FIG. 7B provides peakcurrents measured in three independent runs for the four nucleosides(cross hatched bars). FIG. 7B shows that with both electrodesfunctionalized, a narrow distribution of current peaks is observed at acharacteristic current for each nucleoside. Cross-hatched boxes (3repeated data sets) show peak currents for each nucleoside measured at abaseline current of 6 pA, V=0.5V. The error bars on each box representthe full-width of the measured current distributions. The shaded boxesshow the currents measured when only one of the two electrodes isfunctionalized. Reads for a functionalized surface and a bare Pt (lightshaded bars) and bare Au (dark shaded bars) probe are relativelyinsensitive to the identity of the nucleoside, as shown quantitativelyin FIG. 7C where the junction resistance is plotted vs. the molecularresistance determined with two functionalized probes.

FIG. 8 shows that the read frequency falls as an exemplary tunnel gap ismade larger (i.e., the tunnel current baseline, G_(bl) is made smaller).

FIG. 9 graphically depicts an exemplary embodiment of the presentinvention.

FIGS. 10A and 10B provide details of an exemplary tunnel gap andnanopore array utilizing a gold or titanium nitride probe and a gold ortitanium nitride coated nanopore. The scale bar (110) in the electronmicrograph of FIG. 10B is 2 nm.

FIGS. 10C and 10D provide details of an exemplary tunnel gap andnanopore array utilizing a carbon nanotube probe and a graphenenanopore. The scale bar (210) in the electron micrograph of FIG. 10D is10 nm.

FIG. 10E (cross sectional view) and 10F (top-down view) provide detailsof an exemplary tunnel gap and nanopore array utilizing metal probe andan array of carbon-nanotube nanopores.

FIG. 11 illustrates gap chemistry for an embodiment of the invention.

FIG. 12 illustrates an exemplary embodiment utilizing a gap in a carbonnanotube to form the electrode pair.

FIG. 13 shows exemplary hydrogen bonding sites for various amino acids.

FIG. 14 provides exemplary chemical structures of nucleosides modifiedwith TBDMS.

FIG. 15 provides exemplary current-time traces for G_(bl)=12 pS for 4.3μM dT (FIG. 15A), 2.9 μM dG (FIG. 15B) and 0.8 μM dC (FIG. 15C) in TCBwith both electrodes functionalized with 4-mercaptobenzoic acid. Thecurrent scales for FIGS. 15A and 15B are the same.

FIG. 16 provides exemplary noise spectra in open loop (FIG. 16A) andunder servo control (FIG. 16B) with the gains settings used foracquiring the spike data. Dashed lines are fits to a 1/f spectrum.

FIG. 17 shows two Gaussian log fits to an exemplary data set analyzedwith a fixed (5 pA) cut off (circles), a variable 1.5σ cut-off (squares)and a variable 2σ cut off (triangles). The fitted peak shifts from 6.6to 7.1 pA, a negligible change relative to the separation of peaks fordifferent nucleosides.

FIG. 18 shows exemplary current distributions obtained at G_(bl)=20 pS(V=0.5V) for bare electrodes for dA (FIG. 18A), dC (FIG. 18B), dG (FIG.18C), and dT (FIG. 18D). The total counts recorded in 180 s at anucleoside concentrations listed in Table 1 are listed on each panel.See Example 1.4.

FIG. 19 shows exemplary current distributions obtained at G_(bl)=40 pS(V=0.5V) for bare electrodes for dA (FIG. 19A), dC (FIG. 19B), dG (FIG.19C), and dT (FIG. 19D). Note that some unambiguous reads of dA could bemade because of the width of the distribution. The read rates are lessdisparate between purines and pyrimidines compared to the data acquiredat G_(bl)=20 pS. See FIG. 18. Counts in a 180 s period are listed oneach panel.

FIG. 20 shows an exemplary parabolic fit (solid line) to the data fromFIG. 19A plotted on a log-log plot.

FIG. 21 provides an exemplary distribution for a mix of dT and dC, withthe concentration of dT halved relative to that used for FIG. 5C. Thedistribution is fitted with three Gaussian log functions (solid line).The dT peak is located at 6.6 pA, the dC peak at 12.3 pA. The highcurrent tail is fitted with a small fraction of dG+dC reads (centered at6.6+12.3 pA).

FIGS. 22A and 22B provide exemplary current distributions measured atI_(bl)=6 pA with a bias of 0.75V (G_(bl)=8 pS) resulting in a largergap. The molecular conductances are reduced as expected (G(dA)=14.4 pS,G(dC)=15.6 pS). The reduction for dA is greater than would be predictedby the fit shown in FIG. 7A. This indicates that there is a biasdependence in addition to the exponential dependence on gap size. Datataken as a function of bias at a fixed gap size (See FIG. 23) tend toconfirm this trend.

FIG. 23 provides exemplary peak currents for dA as a function of bias ata constant gap conductance (12 pS; 0.75V, 9 pA, 0.5V, 6 pA and 0.25 V 3pA). The circles are data taken with one bare gold electrode, and theyshow little dependence on bias. Data taken with 2 functionalizedelectrodes (squares) show that peak current increase as bias is lowered.Altering the sign of the bias does not lead to large changes (datum at−0.5 V).

FIG. 24 shows exemplary reads per second with functionalized probesobtained by averaging data obtained over 180 s for dA (circle) and dT(squares) as a function of the baseline tunneling current. These dataare somewhat dependent on probe geometry, an effect reflected in errorbars obtained by comparing data taken with two different probes.

FIG. 25 shows exemplary current distribution (functionalized probes)obtained at G_(bl)=20 pS for dA showing evidence of 2 and even 3molecule reads.

FIG. 26 shows exemplary fitted distributions to sets of experimentaldata (including the 2 molecule read peaks) for dT (circle), dG (square),dC (triangle) and dA (diamond) for I_(b)=6 pA, V=0.5V. (peaks left toright: dT, dG, dC, dA). Setting discrimination levels at the valuesshown (8 pS, 11.7 pS and 14.8 pS) yields a 72% probability for dT if i<8pA, 64% probability for dG (8 pA<i<11.7 pA), 61% for dC (11.7<i<14.8 pA)and 60% for A (i>14.8 pA).

FIG. 27 shows exemplary current distributions obtained by retaining allrecorded spikes in an exemplary analysis (FIG. 27A) and by rejectingspikes of only 1 (20 μs) or 2 (40 μs) sample points duration (FIG. 27B).Data are for dG, 2.9 μM, G_(bl)=12 pS. Solid curved lines are 2 Gaussianlog fits with independent peaks in A, and with peaks fixed at i₀ and 2i₀ in B. The data in A are dominated by a feature at 7.3 pA, equal tothe current recorded with an unfunctionalized probe. The peak in B hasmoved to 9.7 pA. The distribution reflects the finite frequency responseof the STM current-to-voltage converter which will reduces the measuredamplitude of fast peaks (see FIG. 29). Similar results were obtained fordC (peak for all data=7.3 pA, peak for filtered data=13 pA). Data for dTwere not affected by filtering (peak for all data=7.3 pA, peak forfiltered data=6.8 pA).

FIG. 28 provides exemplary current distributions obtained by retainingall recorded spikes in an exemplary analysis for dA (FIG. 28A—all data,FIG. 28B—fastest two points rejected). In this case, a residue of the“single functionalized electrode” peak at 6.5 pA remains when 1 or 2point spikes are rejected, so the data were fitted with three Gaussianlog functions, with two independent peak values (i₀, i₁ and 2i₁). Themaximum of the distribution moves from 6.5 pA to 15.6 pA when theshortest spikes are rejected. The shape of the distribution depends onthe probe used, 2 Gaussian fits working well for other data sets (e.g.,FIG. 2) where the “unfunctionalized” peak was almost completelyeliminated by rejection of the fastest spikes.

FIG. 29 shows exemplary distributions of spike lifetimes for dA (FIG.29A), dC (FIG. 29B), dG (FIG. 29C), and dT (FIG. 29D). Circle lines arefor bare electrodes, triangle lines for one functionalized electrode,and square lines are for two functionalized electrodes. Sharp featuresreflect the data binning. All data are taken at the workingconcentrations shown in Table 1 and 0.5V. G_(bl)=20 pS for bare probesand 12 pS for functionalized probes. The arrows point to the spikes of40 or 20 μs duration that were rejected from the current distributionsto enhance selectivity. With the exception of dT, lifetimes are a littlelonger with two functionalized probes. However, lifetimes for bareprobes, or one functionalized probe are essentially identical. Thus thenarrowing of the tunneling distribution must reflect a difference in therange of allowed bound geometries between functionalized and bare metalsurfaces, and not a difference in bound-state lifetimes. The −3 dBfrequency of the current-to-voltage converter is ˜7 kHz (143 μs) sofaster features are attenuated in the data shown here.

FIG. 30 illustrates cyclic voltammetry for an exemplary bare gold wirein 50 mM potassium ferricyanide (potential vs. Ag wire).

FIG. 31 illustrates cyclic voltammetry for an exemplary HDPE coated STMtip. Assuming a hemispherical exposed tip shape and using the formulaI_(max)=2 πRnFCD, the exposed surface area of coated scanning probes ison the order of 10⁻² μm².

FIG. 32 illustrates exemplary FTIR spectra of 4-mercaptobenzamidemonolayer (lower line) and powder (upper line).

FIG. 33 is an STM image showing islands of mercaptobenzamide on an Ausurface. Image in 1 mM PB buffer with a gold tip, 0.5 V tip bias with 10pA set point.

FIG. 34 shows optical and transmission electron microscope (TEM) imagesfrom exemplary electrodes. FIG. 34A is an optical image of a bareelectrode. FIGS. 34B and 34C are TEM images of a bare electrode. FIG.34D is an optical image of a coated electrode. FIGS. 34E and 34F are TEMimages of coated electrodes. The dashed are in 34C has a radius of 16nm. The arrows in 34E and 34F indicate the location of the exposed gold.

FIG. 35 illustrate telegraph noise in water with a bare electrode probeand a functionalized electrode surface. Similar signals were seen whenboth the probe and surface were bare and also in PB when either surfaceand/or probe was bare.

FIG. 36 shows tunnel current decay curves in pure H₂O (multiple curvesare plotted in each case) for bare gold electrodes (FIG. 36A),functionalized electrodes (FIG. 36B), and one bare and one functionalizeelectrode (FIG. 36C).

FIG. 37 shows exemplary histograms of beta, the negative of the slope ofthe logarithmic decay curves, i.e.,

$- {\frac{{\ln (i)}}{z}.}$

Values are obtained in pure water for bare gold electrodes (FIG. 37A),functionalized electrodes with mercaptobenzamide (FIG. 37B), and onebare and one functionalize electrode with mercaptobenzamide (FIG. 37C).Gausian fits (mean±SD) yield: 37A→6.11±0.68 nm⁻¹, 37B→14.16±3.20 nm⁻¹;and 37C→6.84±0.92 nm⁻¹.

FIG. 38 shows an exemplary 10 s time trace for d(CCACC) taken using4-mercapto benzamide reader molecules. Note the preponderance ofA-signals. The current spike distribution (inset) is almost completelydominated by A-signals with the C component in the fit (black line)being 7% or less. This shows that the probe spends more time bound tothe minority of A bases.

FIG. 39 shows exemplary current spike traces over time for the dAMP(FIG. 39A), dCMP (FIG. 39B), dGMP (FIG. 39C), and dmCMP (FIG. 39D)showing exemplary bursts of data. Each of these examples is surroundedby spike-free regions of current.

FIG. 40 illustrates exemplary current distributions measured forcytidine (grey) and ^(5me)cytidine using benzoic acid readerstrichlorobenzene solvent.

FIG. 41 shows exemplary distribution of “on times” for dGMP, dCMP, dAMP,and d^(m)CMP monomers with the solid lines being exponential fits.

FIG. 42 shows exemplary distribution of “on times” for d(C)₅, d(A)₅, andd(^(m)C)₅ polymers with the solid lines being exponential fits.

FIG. 43 shows exemplary distribution of “off times” for dGMP, dCMP,dAMP, and d^(m)CMP monomers with the solid lines being exponential fits.

FIG. 44 shows exemplary distribution of “off times” for d(C)₅, d(A)₅,and d(^(m)C)₅ polymers with the solid lines being exponential fits.

FIG. 45 shows an exemplary distribution of counts for spikes >0.1 nA ford(A)₅. These are about 20% of the total and are not observed in dNTPs ord(C)₅.

FIG. 46 shows an exemplary distribution of counts for spikes >0.1 nA ford(^(m)C)₅. These are about 20% of the total and are not observed indNTPs or d(C)₅.

FIG. 47 shows exemplary SPR sensorgrams of nucleoside-5′-monophosphates(A, C, G, T, R) interacting with the benzamide surface (R: 2-Deoxoribose5-phosphate sodium salt containing no DNA base). The lines are fittedcurves modeled to describe a 1:1 binding event.

FIG. 48 shows histograms of adhesion events, as recorded with an atomicforce microscope, for a pair of 4-mercaptbenzamide reader moleculestrapping dAMP. FIG. 48A shows a control taken in the absence of dAMP,which showed almost no adhesion events between the benzamide molecules,presumably because they were blocked by water. FIG. 48B shows theadhesion of dAMP after a first rinse. FIG. 48C shows adhesion of dAMPafter a second rinse. FIG. 48D shows adhesion of dAMP after a thirdrinse. FIG. 48E shows adhesion of dAMP after a fourth rinse. Addition ofdAMP led to a number of adhesion events that increase as excess dAMP wasrinsed out of the system (FIGS. 48B,C) and then decreased as the rinsingcontinued (FIGS. 48D, E).

FIG. 49 shows exemplary simulated displacement (top pattern—black) andcurrent (bottom pattern—grey) vs. time-steps where the correlation, C,has a value of 0.9.

FIG. 50 shows exemplary simulated displacement (top pattern—black) andcurrent (bottom pattern—grey) vs. time-steps where the correlation, C,has a value of 0.98.

FIG. 51 shows exemplary simulated displacement (top pattern—black) andcurrent (bottom pattern—grey) vs. time-steps where the correlation, C,has a value of 0.99.

FIG. 52 shows exemplary normalized distributions for signals obtainedfrom homopolymers. FIG. 52A shows fits to normalized currentdistributions. FIG. 52B shows normalized spike frequencies in a signalburst, fitted with polynomials. The fits to the distributions are usedto assign the probability that a particular noise burse originates froman A or a C (if the average currents and frequencies lie above or belowthe crossover points labeled “I_(AC)” and “f_(AC)”). Currentdistributions for C and ^(m)C are separated (crossover point labeled“I_(mC)”) but frequency distributions overlap.

FIG. 53 shows exemplary hydrogen bonding modes of 4-mercaptobenzamidefor adenine, thymine, cytosine, and guanine.

FIG. 54 shows exemplary data from reading a single base within aheteropolymer. 54B shows exemplary bursts of tunneling noise with large,infrequent spikes signaling C and smaller, more frequent spikessignaling A. The “*” labeled spike is nonspecific. FIG. 54C shows therolling average of the spike height (0.25 s window, 0.125 s steps). Cbases generate a negligible number of spikes below 0.015 nA (straightline). FIG. 54D shows the rolling average of the spike frequency. FIG.54E shows the probability that the signal comes from an A (shown by thelight grey line) or a C (shown by the darker line).

FIG. 55 shows tunneling signals for a functionalized tunnel gap in aphosphate buffered saline, but without analyte, with a 20 pS gap (i=10pA, V=+0.5V). This example gave a signal free of features, except forsome AC coupled line-noise pointed to by arrows.

FIG. 56 shows tunneling signals for a functionalized tunnel gap. FIGS.56C-F show characteristic current spikes produced when nucleotides dAMP(FIG. 56C), dCMP (FIG. 56D), d^(m)CMP (FIG. 56E), and dGMP (FIG. 56F)were introduced (dTMP gave no signals in this example). FIGS. 56G-J showcorresponding distribution of pulse heights for dAMP (FIG. 56G), dCMP(FIG. 56H), d^(m)CMP (FIG. 56I), and dGMP (FIG. 56J). The line curves inFIGS. 56G-J are fits to two Gaussian distributions in the logarithm ofcurrent.

FIG. 57 illustrates parameters used to characterize the tunnelingsignals. Spikes are counted if they exceed a threshold equal to 1.5× thestandard deviation of the noise on the local background. The signalsoccur in bursts (duration T_(B), frequency f_(B)) each containingcurrent spikes at a frequency f_(S). The spikes stay high for a periodt_(on) and low for a period t_(off). The total count rate (see FIGS.56G-J) is the number of spikes in all bursts divided by the measurementof time.

FIG. 58 illustrates how tunneling signal distributions from oligomersresemble those of the constituent nucleotides. FIGS. 58A, C, and E showrepresentative current traces from d(A)₅, d(C)₅, and d(^(m)C)₅ with thecorresponding distributions shown in FIGS. 58B, D, and F. The linecurves in FIGS. 58B, D, and F show fits for both constituent nucleotidesand oligomers nucleotides. The line curves correspond closely with oneanother. FIGS. 58G and I show current traces from mixed oligomersd(ACACA) (FIG. 58G) and d(C^(m)CC^(m)CC) (FIG. 58I) with correspondingcurrent distributions (FIGS. 58H and J). The solid line curves in FIGS.58H and J are scaled homopolymer fits. In FIG. 58H, the top dashed curveshows the A contribution and the bottom dashed curve shows the Ccontribution. In FIG. 58J, the top curve shows the ^(m)C contributionand the bottom curve (beginning at 0.02 on the Current (nA) axis) showsthe C contribution. The data are well described by the homopolymerparameters though some intermediate signals (labeled as “1”) and newhigh current features (labeled as “2”) show that the sequence contextaffects the reads slightly. The upper bar in FIG. 58G marks C-likesignals, while the lower bars mark A-like signals. The upper bars inFIG. 58I mark C like signals and the lower bars mark ^(m)C-like signals.

FIG. 59 shows data regarding the lifetime of the reading complex (thelifetime is on the order of a second at zero force). FIG. 59A shows AFMgap functionalization where the s-shaped line represents a 34 nm PEGlinker. FIG. 59B shows representative force curves showing (i) pullingon more than one molecule at a time—the force baseline is not restoredafter each break and the z-extension (corrected tip displacement) is ˜34nm and (ii) a single molecule curve of the type accepted by exemplarysoftware (as described by A. Fuhrmann, PhD Thesis in Physics, ArizonaState University, 2010). The force returns to the baseline after thebond breaks and the corrected extension is ˜34 nm. FIG. 59C showshistograms of bond breaking forces at the pulling speeds marked. Thecurves show exemplary maximum likelihood fits to the heterogeneous bondmodel. FIG. 59D shows bond survival probability plotted versus bondmodel parameters (solid lines) which are, from top to bottom 5000 nm/s,2000 nm/s, 500 nm/s, and 200 nm/s. These fits yield a zero-force offrate of 0.28 s⁻¹ implying that the assembly lives for times on the orderof seconds in a nanogap, much longer than the lifetime in solution (seeFIG. 47 for details of solution binding measurements).

FIG. 60 illustrates a chip according to an embodiment of the invention.In the chip, a pair of reading electrodes are formed by atomic layerdeposition of conductors, such as TiN separated by a thin (for example,2 nm) dielectric layer with nanopores drilled through the chip.Recognition molecules (reagents) are covalently tethered to the metalelectrodes and form a self-assembled junction on each base (or residue)in turn via non-covalent interactions as electrophoresis drives themolecule through the gap.

FIG. 61 shows exemplary hydrogen bonding modes ofimidazole-2-carboxyamide for adenine, thymine, cytosine, and guanine.

FIGS. 62A and 62B show exemplary current distributions measured withdeoxy-nucleotides in a fixed gap comprised of imidazole-2-carboxyamidefunctionalized electrodes.

FIG. 63 shows an exemplary device with imidazole-2-carboxyamidefunctionalized electrodes and a probe that may be translated across asurface at a constant gap.

FIG. 64 shows that the DNA reads fit nucleotide current distributionsfor dC₅ (FIG. 64A) and for dA₅ (FIG. 64B).

FIG. 65 shows exemplary current reads for an AAAAA oligomer for a fixedgap.

FIG. 66 shows exemplary current reads for a CCCCC oligomer for a fixedgap.

FIG. 67 shows exemplary current reads for a d(^(m)C)₅ oligomer for afixed gap.

FIG. 68 shows exemplary current reads for a d(CCCCC) oligomer for avariable gap held at an approximately constant value by servo control ofthe tunnel current.

FIG. 69 shows an exemplary plot of signal burst time vs. reciprocal scanspeed for dA₅ (top slope) and dC₅ (bottom slope) with a slope ofapproximately 0.3 nm.

FIG. 70 shows exemplary current reads for an ACACA oligomer for a fixedgap.

FIG. 71 shows exemplary current reads for an CCACC oligomer for a fixedgap.

FIG. 72 shows exemplary current reads for an C^(m)CC^(m)CC oligomer fora fixed gap.

FIG. 73 shows exemplary current reads for an ACACA oligomer for avariable gap held at an approximately constant value by servo control ofthe tunnel current.

FIG. 74 shows exemplary current reads for an C^(m)CC^(m)CC oligomer fora variable gap held at an approximately constant value by servo controlof the tunnel current.

FIG. 75 shows exemplary current reads for an GTCGTCGTC oligomer for avariable gap held at an approximately constant value by servo control ofthe tunnel current.

FIG. 76 shows exemplary recognition molecules (reagents) for amino acidsand the peptide backbone.

DETAILED DESCRIPTION OF THE INVENTION

The present invention provides compositions, components, devices, andmethods for analyzing polymer units and/or polymers. Exemplary polymerunits and polymers that may be analyzed include heteropolymers andassociated units. For example, polymers that may be analyzed includeDNA, RNA, polysaccharides, and peptides; and polymer units includepolymer monomers, nucleotides, nucleosides, amino acids, polysaccharidesmonomers. In some embodiments, epigenetic marks, such as methylated DNAand/or RNA, may be analyzed and distinguished from, for example,non-methylated DNA/RNA units.

The devices include two or more electrodes functionalized with one ormore reader molecules (also referred to as reagents) and a tunnel gapthrough which the polymer units and/or polymer may pass. The reagents onthe electrodes are capable of forming a transient bond to the units ofthe polymer. A transient chemical or physical bond forms when the unitis in the gap and completes the circuit between the first and secondelectrodes. The formed transient bond may then elicit a detectablesignal that is used to analyze the polymer.

Electrodes

The two or more electrodes may be made of any suitable material that canbe functionalized with a reagent capable of binding the target polymerunit(s). For example, the electrodes may be made of any conductivematerial, such as a metal, a metal alloy, gold, platinum, a gold alloy,a platinum alloy, carbon, carbon nanotubes, graphene, or titaniumnitride. In some embodiments, the electrodes comprise a probe and asubstrate. The electrodes may be formed on or in between or be partiallyinsulated with any suitable inorganic or organic insulating material,such as inorganic materials including Si_(x)O_(-1x), silicon nitride,metal oxides, or organic materials, including polymers such aspolyethylene, polystyrene, polymethylmethacrylate and others as are wellknown in the art. The insulating material may be configured to preventbackground noise from the electrode when a current is flowing. Forexample, an electrode may be completely covered with HDPE except for asmall tip or apex. In another embodiment, the electrode may be embeddedbetween insulating layers with only regions in contact with the nanoporeexposed (FIG. 60). As much as a square micron can be exposed to saltsolutions of up to one molar with negligible leakage currents for biasesof half a volt or less.

The reading reagents play an important role in “sharpening” theelectrodes. The typical gold electrode has a nanocrystalline compositionin which facets of 10 nm or more in size are exposed. Thus it wouldappear to be impossible to contact just a single base. However, singlebases are readily resolved when the electrodes are functionalized. Thisis because specific molecular contacts now serve as the tunnelingelectrodes, forming sharp, well defined asperities on the metal surface.

Reagents

The electrodes are functionalized by one or more reagents. Theelectrodes may be functionalized with the same reagent, a combination ofreagents, or individually functionalized with separate reagents. Anysuitable reagent capable of binding to the electrode and transientlybinding to the target polymer unit may be used.

To facilitate binding of the target polymer unit to the electrode, avariety of functional groups may be tethered to a reader moleculecapable of binding to the target polymer units, depending on theelectrode substance desired. Suitable functional groups may include, forexample, —SH, —NH₂, —N₃, —NHNH₂, —ONH₂, —COOH, —CHO, acetylene,dithiocarbamate, and dithiocarboxylate. Dithiocarbamate linkage to ametal greatly increases the tunnel current by aligning molecular levelsmore closely with the metallic Fermi level (see Florian von Wrochem,Deqing Gao, Frank Scholz, Heinz-Georg Nothofer, Gabriele Nelles andJurina M. Wessels, “Efficient electronic coupling and improved stabilitywith dithiocarbamate-based molecular junctions”, Nature Nanotechnology,Jun. 20, 2010.) In some embodiments, the electrode is tethered to afunctional group which can then bind to the reader molecule. Forexample, with metals, when the electrode is gold, a reagent with a thiolfunctional group may be used to facilitate a covalent bond between thereagent and the electrode. Dithiocarbamates may be used to bind to gold,Pt, and TiN. These groups can provide enhanced electronic couplingbetween the metal and the reagent. Amine chemistry may be used tofunctionalize graphene pores and carbon nanotubes ends since grapheneedges frequently have carboxylates, carbonyls, and epoxides.

The reagents are capable of forming a transient bond with the targetpolymer unit (J. He et al., Nanotechnology 20, 075102 (2009)) and thenucleosides (S. Chang et al., Nature Nanotechnology 4, 297 (2009)). Thetransient bond may be a physical, chemical, or ionic bond so long as thebond permits a detectable electronic signal to be detected via theelectrodes (S. Chang et al., Nanotechnology 20, 075102 (2009); M. H.Lee, O. F. Sankey, Phys. Rev. E 79, 051911 1 (2009)). A preferredtransient bond includes a hydrogen bond. As such, exemplary reagents mayinclude hydrogen donating or accepting groups. Another embodiment is thepi-stacking interaction between aromatic rings that are pushed togetherin water or aqueous electrolytes.

Exemplary reagents for binding DNA and/or RNA include mercaptobenzoicacid, 4-mercaptobenzamide, imidazole-2-carboxide, anddithiocarbamateimidazole-2-carboxide (also referred to as4-carbamonylphenyldithiocarbamate). 4-mercaptobenzamide presents twohydrogen-bond donor sites (on the nitrogen) and one hydrogen-bondacceptor site (the carbonyl). Likely binding modes to, for example, thefour nucleotide bases are shown in FIG. 53. Likely binding modes ofimidazole-2-carboxyamide to the four nucleotide bases are shown in FIG.61. Additional bonding modes that involve pi-stacking between thearomatic rings in the readers and DNA bases are also likely. Thereagents may be formulated to present one or more hydrogen bond donorsand/or one or more hydrogen bond acceptors when in various solvents,such as organic solvent, water, or an aqueous electrolyte solution. Forexample, mercaptobenzoic acid works in organic solvent, such astrichlorobenzene, while 4-mercatobenzamide, imidazole-2-carboxide, anddithiocarbamateimidazole-2-carboxide work in aqueous electrolytes andwater. It should be noted that many molecules that embody these designprinciples will function as readers. For example, guanine,functionalized with a thiol to anchor it to gold or TiN electrodes, willgenerate recognition signals.

In some embodiments, the reagents may be configured to include aflexible moiety that forms a bridge between the electrode and hydrogenbonding moiety of the reagent. This bridge may be a substituted orunsubstituted alkyl chain, such as —(CH₂)_(y)—, where y is an integer of1 to 5. For example, when functionalized to the electrode,imidazole-2-carboxyamide has a —CH₂CH₂— bridge connecting the amideportion to the electrode. This bridge permits the amide portion torotate and, thereby, interact in different and detectable ways withadenine, cytosine, guanine, and thymine. See FIG. 61.

Reagents may also be configured to form transient bonds with amino acidsto analyze peptides. FIG. 13 shows examples of hydrogen bond donor andacceptor sites on amino acids. Two or more nearby sites (labeled “D”,donor or “A”, acceptor) are available on asparagine, glutamic acid,glutamine, histadine and arginine. Single sites on lysine, serine,threonine, tyrosine and tryptophan could be read in conjunction using areagent that also forms hydrogen bonds to the peptide backbone. Aromaticreagents can recognize amino acids with aromatic rings (histadine,tyrosine, proline and tryptophan) by means of pi-stacking. Accordingly,peptides may be analyzed according to the devices and methods describedherein. FIG. 76 shows exemplary reagents for recognizing a peptidebackbone and for recognition of side chains of amino acids.

Tunnel Gap

Units of a polymer, such as nucleosides of DNA or amino acids of aprotein are detected as they diffuse through a tunneling gap or aredriven through it by electrophoresis. The width of the gap may be fixedor dynamically adjustable. The gap is preferably fixed. The gapcomprises the space between the two electrodes. The gap is adjusted to asize such that each target unit fits into the gap. The gap may have awidth of about 0.5 to about 6 nm, such as about 1 to about 4, about 1.5to about 3.5 nm, about 2 to about 3 nm, or about 2 to about 2.5 nm. Thegap width may vary depending on the reagent being used and the targetpolymer unit(s) to be analyzed. In the case of two electrodesfunctionalized with 4-mercaptobenzamide, the gap may be about 2 to about2.5 nm, such as about 2.1 to about 2.2 nm, or about 2.16 nm (when thegap conductance is 20 pS—typically used for DNA reads). In the case oftwo electrodes functionalized with imidazole-2-carboxyamide, the gap maybe from about 2.2 to about 2.6, such as about 2.3 to about 2.5 nm; fromabout 2.35 to about 2.4 nm, or about 2.37 nm (when the gap conductanceis 20 pS—typically used for DNA reads). The gap distance may bedetermined as described below. FIG. 1 shows the distinctive hydrogenbonding formed with each of the four DNA nucleosides in one fixed tunnelgap. The 4-mercaptobenzene forms hydrogen bonds with each of the fournucleosides. Hydrogen bonds are circled and “S” stands for thedeoxyribose sugar moiety. These structures were generated by computersimulation, and probably represent the actual structure quite well,because the 4-mercaptobenzamide was used in organic solvent. In the caseof the reader molecules that work in water, attempts to model thestructures are complicated by competition for hydrogen bonds with watermolecules, and the interactions of aromatic rings mediated by waterwhich pushes them together to form pi-stacking interactions.

The exact size of the gap may be important in obtaining reliable reads.It should be sized so that the majority of time (or majority of signalsgenerated) are caused by the presence of just one unit of the polymer inthe gap. Suitable gap widths may be determined by using a device capableof a dynamically adjusting the gap. In some embodiments, a dynamicallyadjustable device may be used to analyze target units. In either case,the gap width may be determined or set as follows: The electrodes areapproached together until a chosen tunnel current is achieved at aparticular bias. For example, a current of 6 pA at 0.5V bias correspondsto a gap of 2.5 nm when tunneling in 1,2,4-trichlorobenzene. The gap ismaintained by applying active servo control as is well known in the artfor scanning tunneling microscopy. In certain embodiments, the servocontrol has a frequency of response limited to 100 Hz or less.

Provided that the gap is kept large enough, the background tunnelingsignal is free of features, as shown in FIG. 2. When molecules bind inthe gap, transient current spikes are observed in the tunnel gap (FIG.4). With both electrodes functionalized, a narrow distribution ofcurrent peaks is observed at a characteristic current for eachnucleoside (FIG. 7B). However, identifying the specific nucleoside fromthe current signal is complicated by a high-current tail on thedistribution, attributable to more than one target molecule binding inthe gap simultaneously. This may give rise to a second peak in thecurrent distribution at twice the current of the main peak (FIG. 6). Therelative frequency of the multi-molecule reads increases as the gap ismade smaller (baseline conductance or current increased) so that morecontacts across the electrodes become possible. This increasingfrequency of two molecule reads is shown in FIG. 7A (which also showshow the peak currents increase with decreasing gap). Thus, making thegap larger has two deleterious effects: first, the peak currents fall(as shown in FIG. 7A), and second, the read-rates decrease for a givenconcentration of target (FIG. 8).

Thus, in some embodiments, important elements include (a) theincorporation of a variable and controllable tunnel gap with a nanoporethrough which the target polymer can be translocated to present oneelement at a time to the reading system (i.e., one base at a time for aDNA polymer); and (b) the use of reagents that bind all targets in somemanner or another, the gap being adjusted to such a size that just a fewdistinct binding geometries exist for the targets, thereby generatingdistinctive signals.

Nanopores

In some embodiments, the device may be equipped with one or morenanopores, through which the polymer may be directed to the tunnel gapfor analysis. The nanopores may be configured to permit the flow of thepolymer to the tunnel gap one unit at a time. Thus, a nanopore foranalyzing DNA may be smaller than a nanopore for analyzing a peptide.

Such an embodiment is illustrated in FIG. 9 for a tunnel junction with avariable gap. Details of the gap itself are given in FIGS. 10A-F.Details of a fixed gap device are given in FIG. 60, A polymer to besequenced, such as DNA, is present in a fluid reservoir 1 (shown incross section). The fluid containing the polymer may flow through anarray of nanopores 3, or may be optionally driven through the nanoporesby an electrophoretic bias, V_(e) applied between the first reservoir 1and a second fluid reservoir 2 by means of reference electrodes 4. Bothreservoirs are filled with electrolyte, for example 1M KCl, and, inaddition, the pH of reservoir 1 may be adjusted to a large value (pH=11or 12) in order to maintain the target DNA in its single stranded form.If independent verification of translocation from electrical signals isnot required, the electrolyte solution in the collection reservoir 2 ispreferably made small (mM) to minimize electrochemical leakage. Incertain embodiments, the nanopores are electrically conductive andconnected by an electrode 5 plated onto the top of the nanopore array.The array may be probed by one or more second electrodes 6 held in placeby a scanning transducer 7, such as the x,y,z scanning elements wellknown in the art of scanning probe microscopy. The scanner may beattached to the nanopore array via the rigid frame 8.

Exemplary illustrations of the tunnel junction are shown in FIG. 10.FIG. 10A shows a gold probe reading sequence as a DNA molecule passesbetween the probe 104 and a gold or TiN electrode 103 on top of ananpore 101 through which the DNA is translocated by electrophoresis.The nanopore 101 is shown in cross section—it is drilled through asilicon, silicon nitride or silicon dioxide substrate 102. Readingreagents 105 and 106 are attached to the probe 104 and metal electrode103. An electron micrograph in FIG. 10B shows a nanopore 107 in asilicon nitride substrate that has been coated with a thin (20 nm) layerof gold 108. The act of drilling the pore causes the gold torecrytaliize around the pore, so that a sharp, atomically ordered ledgeof gold 109 protrudes over the edge of the nanopore to form one of thereading electrodes for the polymer.

FIG. 10C shows an embodiment in which a probe bearing a carbon nanotubeelectrode 204 is held over a nanopore 201 in a graphene substrate 202supported on a silicon nitride substrate 203. Reading reagents 205 and206 are attached to the CNT 204 and edges of the graphene nanopore 201.The electron micrograph in FIG. 10D shows a nanopore 207 drilled in agraphene multilayer 208.

FIGS. 10E and 10F show an embodiment in which a metal probe 13protruding from insulation 12 is held over an array of carbon nanotubenanopores (one nanopore is labeled as 10) protruding through adielectric substrate 11 coated with a thin metal electrode 5 throughwhich the carbon nanotubes protrude. Such an array may be fabricated byCVD growth of nanotubes from a silicon surface, subsequently filled withsilicon nitride with the underlying support etched away, as described byHolt et al., Fast Mass Transport Through Sub-2-Nanometer CarbonNanotubes. Science, 2006. 312: p. 1034-1037, hereby incorporated byreference. For example, the top side of the membrane may have a layer ofAu evaporated onto it to act as a contact (of thickness 10 to 100 nm)before the remaining protruding carbon nanotubes are removed by plasmaetching. Electrophoretic translocation of DNA through carbon nanotubeshas recently been demonstrated by Liu et al., Translocation ofsingle-stranded DNA through single-walled carbon nanotubes. Science,2010, 327, p 64-67, hereby incorporated by reference. The probe 6 maybecovered with a layer of insulation 12 leaving a small amount (few squaremicrons) of the apex exposed 13 as described by Nagahara et al.,Preparation and Characterization of STM Tips for ElectrochemicalStudies. Rev. Sci. Instrum. , 1989. 60: p. 3128-3130. This results inminimal electrochemical leakage current into the electrolyte solution.

In some embodiments, the components described above may be configuredinto a microarray or chip as illustrated, for example, in FIG. 60. Herea supporting material 302 (silicon, silicon oxide or silicon nitride) iscoated with a thin metal electrode 303, for example TiN deposited byatomic layer deposition, then covered with a layer of a dielectric 304of a thickness 308 chosen to be optimal for the reading reagents used.For most of the reagents described here this thickness is from 1.5 to 3nm. A second electrode 305 is deposited and covered with a finaldielectric layer 309. A nanopore 301 is drilled through the entiredevice (by means of an electron beam as is well known in the art) andthe exposed metal electrode surfaces functionalized with readingreagents 306 and 307.

Metal electrodes may be formed around a nanopore by any conventionalmethod. For example, Pt may be deposited by Focused Ion Beam chemicalvapor deposition on a nanopore formed in a silicon nitride membrane.Metals, for example TiN, may also be deposited by atomic layerdeposition or chemical vapor deposition and a pore etched or createdthereafter. Metals may also be first coated on a membrane, such as SiN,Si, or SiO₂, and thereafter a pore drilled.

Graphene, an intrinsically conducting substance may also be used to asan electrode with a nanopore. In the case of graphene, a pore may bedrilled in the graphene. Translocation for a graphene pore may be usedfor long oligomers, for example, for up to 48 kbp. When using grapheneit is possible to only functionalize the edges of the pore.

An exemplary tunnel gap that uses a carbon nanotubes for a nanopore isshown in FIG. 11. The probe may be removed from the substrate, sofunctionalization with a different reagent on the substrate and probe ispossible. In certain embodiments, the probe, gold or platinum, or aplatinum alloy, is functionalized with 4-mercaptobenzcarbamide 21, areagent that presents a hydrogen bond donor and acceptor in aqueoussolution. The end of the carbon nanotube 10 may preferably befunctionalized with carbamide moieties 22 using amide linkages (notshown) as is well known in the art. See Feldman, A. K., M. L.Steigerwald, X. Guo, and C. Nuckolls, Molecular Electronic Devices Basedon Single-Walled Carbon Nanotube Electrodes. Acc. Chem. Res., 2008. 41:p. 1731-1741.

Control of Translocation Speed

An additional advantage of using a reading gap functionalized withreadding reagents is the long intrinsic binding time of the bases in thegap, as discussed in Example 13. A major problem with nanopores is thehigh speed of DNA translocation when a bias, big enough to dominatethermal fluctuations, is applied across the pore. The DNA translocatesat speed of millions of bases per second, too fast for any proacticalreadout scheme. This problem is discussed by Branton et al., NatureBiotechnology volume 28, pp 1146-1153, 2008. In the case where theelectrodes are funtionalized with molecules that bind the DNA base, onebase can remain trapped for up to several seconds. A detailed analysisof the atomic force microscopy data presented in FIG. 59 (Huang et al,Nature Nanotechnology, vol 5 pp 868-873, 2010) shows that only a smallforce needs to be applied to increase the speed of the DNA through thenanopore functionalized with reading reagents. For example, the data ofFIG. 59 can be used to show that at a bias of 80 mV across the nanpore,the DNA will translocate at a speed of 10 bases per second, increasingto over 100 bases per second at 120 mV.

Analysis of Polymers

The compounds, components, devices, and methods of the present inventionmay be used to analyze polymers. In certain methods of operation, abias, such as from about 0.1 to 1V, such as from about 0.3 to 0.7V, orabout 0.5V (V_(t)), may be applied between the electrodes by means ofvoltage source, V_(t) in FIG. 9. The gap between the probe and thenanopore is adjusted by a transducer (7 in FIG. 9) until the desiredset-point current is achieved. This may be from about 1 and 10 pA, suchas from about 3 to 6 pA. For example, a translocation bias (V_(e)) maybe applied to generate translocation of DNA through carbon nanotubes.Preferred values of V_(e) are between 0.1 and 1V.

In some embodiments, one of the electrodes (such as a probe) may bemoved over the surface using the lateral scanning motion of thetransducer 7 to locate a nanopore that is successfully translocatingDNA, and the gap adjusted to achieve maximum discrimination in thetunneling signals. The gap would be set to a preferred initial value(e.g., 6 pA current at 0.5V) and small adjustments in background tunnelcurrent made to optimize the separation of the signals from the fournucleosides.

Preferable additional components in analyzing a polymer are: (a)Rejection of fast data spikes (below 40 μs duration); (b) Automatic peakdetection with a threshold set at 1 to 2 standard deviations above thenoise level in a 0.3 s block of data; and (c) Adjustment of the servogain that maintains the background current signal so that the frequencyresponse is not faster than 35 Hz; (d) Means to turn the servo offduring acquisition of data. When the servo that controls the average gapsize is left on during data acquisition, the data are distorted as theservo adjusts the gap in response to the desired sequencing signal. Forexample, the traces in FIGS. 68, 73 and 74 were obtained with the servoon (with a slow response time as just described). The traces in FIGS.70, 71 and 72 were obtained with a fixed gap (no servo control). Thesignals are much easier to intepret. In instruments with a variable gap,an optimal arrangement is to arrange to sample the background signal inthe absence of DNA, stabilize the gap with a servo, and then turn theservo off to acquire data, resetting the gap then the DNA signal ceases.

In certain embodiments, the current signals are selected based on theirduration, and the background current is fitted numerically overintervals of 0.5 s or less so as to establish a baseline for recognizingpeaks above this background.

It will be recognized that a further advantage of the present inventionis that any target that presents hydrogen bond donors and or acceptors(and/or aromatic rings that can pi-stack) that are located a smalldistance (up to 10 carbon-carbon bonds in distance) apart may be read inthis scheme, adjusting the gap, if necessary, to optimize the signalfrom a subset of targets.

A Carbon Nanotube Forming Both Electrodes

In other embodiments a carbon nanotubes may be used to form bothelectrodes as illustrated in FIG. 12. Carbon nanotubes as electrodes forsequencing polymers is described in 61/083,993 (“Carbon Nanotube BasedDevice for Sequencing Polymers”), which is herein incorporated byreference. DNA translocates across a small gap in the carbon nanotube 30(see FIG. 12) using a device built on the surface of a silicon wafer asdescribed in Liu et al., Translocation of single-stranded DNA throughsingle-walled carbon nanotubes. Science, 2010, 327, p 64-67, and furtherin 61/083,993 (“Carbon Nanotube Based Device for Sequencing Polymers”).A very small nick in the CNT may be made by brief exposure of the openpart of the resist barrier 35 to an oxygen plasma etch followed bybending the device with a transducer 36 that pushes against a thinmembrane 34 that flexes relative to fixed points 37 above the gap anddisplaced to one side of it. Bending the nicked carbon nanotube willbreak it, the extent of the gap being increased as the base on which thetube sits is further bent. The size of the opened gap is measured bymeans of a tunnel current that passes from one electrode (31 a) toanother (31 b). When a desired gap size (2 to 2.5 nm) is attained, theends of the CNT are functionalized with carbamide groups 32 as describedabove.

In certain methods, DNA is translocated through the gap 30 and thetransducer 36 adjusted to optimize the separation of the tunnel currentsignals from bases that span the gap by binding carbamide groups onelectrode 31 a and 31 b.

EXAMPLES Example 1 Synthesis and Characterization of Materials 1.1Materials and Methods

Proton NMR (¹H) spectra were recorded on a Varian 500 MHz spectrometer.¹H chemical shifts in chloroform were referenced to the solvent peak(δ_(H)=7.26 ppm). High resolution mass spectra (HRMS) were recordedusing the atmospheric pressure chemical ionization (APCI) technique. TheUV absorbance was recorded on a Varian Cary 300 UV spectrophotometer.Flash chromatography was performed using automated flash chromatography(Teledyne Isco, Inc. CombiFlash Rf). All chemical reagents werepurchased from commercial suppliers and used as received unlessotherwise noted. 2′-Deoxyadenosine and 2′-deoxyguanosine were purchasedfrom TCI America; thymidine from Alfa Aesar; 2′-deoxycytidine fromSigma-Aldrich. Anhydrous N,N-dimethylformamide (DMF) in a Sure/Seal™bottle was purchased from Sigma-Aldrich. 1,2,4-trichlorobenzene (TCB,99%, Aldrich) was dried over molecular sieve (4 Å) under nitrogen, andthen distilled under reduced pressure after filtration. All othersolvents were used as received.

1.2 General Procedure for the Preparation ofbis(tert-butyldimethylsilyl) (TBDMS) Derivatives of Nucleosides (SeeFIG. 14)

See (D. A. Barawkar, R. K. Kumar, K. N. Ganesh, Tetrahedron Letters 48,8505 (1992); W. Zhang, R. Rieger, C. Iden, F. Johnson, Chem. Res.Toxicol. 8, 148 (1996); P. Potier, A. Abdennaji, J. P. Behr, Chem. Eur.J. 6, 4188 (2000).

tert-butyldimethylsilyl chloride (TBDMSCl, 2.5 mmol) was added to asolution of dry nucleoside (1.0 mmol), dimethyl aminopridine (DMAP, 0.15mmol) and imidazole (6 mmol) in anhydrous DMF (10 mL). After thereaction mixture was stirred overnight at room temperature undernitrogen, it was quenched with sat. aq. NaHCO₃, and extracted withdichloromethane. The combined organic layer was concentrated, and theresidue was purified by silica gel flash chromatography with a gradienteluent of CH₂Cl₂—CH₃OH from 100:0 to 100:5.

3′,5′-Bis-O-(tert-butyldimethylsilyl)-deoxyadenosine (1): yield 80%. ¹HNMR (500 MHz, CDCl₃): δ 8.29 (s, 1H, 2-H), 8.09 (s, 1H, 8-H), 6.65 (brs, 2H, NH₂), 6.41 (t, 1H, 1′-H), 4.56 (dd, 1H, 3′-H), 3.96 (d, 1H,4′-H), 3.82 (dd, 1H, 5′-H), 3.72 (dd, 1H, 5″-H), 2.59 (m, 1H, 2′-H),2.39 (m, 1H, 2″-H), 0.86 (s, 18H, (CH₃)₃CSi), 0.05 (s, 6H, CH₃SiO), 0.03(s, 6H, CH₃SiO). HRMS (APCI): calcd for C₂₂H₄₁N₅O₃Si₂+H, 480.2826;found, 480.2818.

3′,5′-Bis-O-(tert-butyldimethylsilyl)-deoxycytidine (2): yield 17%. ¹HNMR (500 MHz, CDCl₃) δ 8.07 (d, 1H, 6-H), 7.14 (br s, 2H, NH₂), 6.24 (t,1H, 1′-H), 5.84 (d, 1H, 5-H), 4.38 (m, 1H, 3′-H), 3.92 (m, 2H, 5′-H),3.77 (m, 1H, 4′-H), 2.42 (m, 1H, 2′-H), 2.08 (m, 1H, 2″-H), 0.92 (s, 9H,(CH₃)₃CSi), 0.88 (s, 9H, (CH₃)₃CSi)), 0.11 (s, 3H, CH₃SiO) 0.10 (s, 3H,CH₃SiO), 0.07 (s, 3H, CH₃SiO) 0.06 (s, 3H, CH₃SiO). HRMS (APCI): calcdfor C₂₁H₄₁N₃O₄Si₂+H, 456.2714; found, 456.2722.

3′,5′-Bis-O-(tent-butyldimethylsilyl)-deoxyguanosine (3): The crudeproduct from chromatography was further purified by recrystallizing inethanol (95%). yield 21%. ¹H NMR (500 MHz, CDCl₃) δ 13.10 (br s, 1H,NH), 7.89 (s, 1H, 8-H), 7.11 (br s, 2H, NH₂), 6.26 (t, 1H, 1′-H), 4.57(t, 1H, 3′-H), 3.97 (t, 1H, 4′-H), 3.81 (m, 1H, 5′-H), 3.77 (m, 1H,5″-H), 2.51 (m, 1H, 2′-H), 2.37 (m, 1H, 2″-H), 0.91 (s, 9H, (CH₃)₃CSi),0.90 (s, 9H, (CH₃)₃CSi)), 0.10 (s, 6H, CH₃SiO) 0.07 (s, 6H, CH₃SiO).HRMS: (APCI) calcd for C₂₂H₄₁N₅O₄Si₂+H, 496.2775; found, 496.2767.

3′,5′-Bis-O-(tert-butyldimethylsilyl)-thymidine (4): yield 83%. ¹H NMR(500 MHz, CDCl₃) δ 9.78 (br s, 1H, NH), 7.40 (s, 1H, 6-H), 6.27 (t, 1H,1′-H), 4.33 (t, 1H, 3′-H), 3.85 (t, 1H, 4′-H), 3.80 (dd, 1H, 5′-H), 3.69(dd, 1H, 5″-H), 2.18 (m, 1H, 2′-H), 1.93 (m, 1H, 2″-H), 1.84 (s, 3H,5-CH₃), 0.85 (s, 9H, (CH₃)₃CSi), 0.82 (s, 9H, (CH₃)₃CSi)), 0.04 (s, 6H,CH₃SiO) 0.00 (s, 6H, CH₃SiO). HRMS: (APCI) calcd for C₂₂H₄₂N₂O₅Si₂+H,471.2711; found, 471.2712.

1.3 Preparation of Stock Solutions

Saturated solutions of nucleosides (dA, dG, dT, dC) with hydroxyl groupsprotected by tert-butyldimethylsiyl groups (1.0 mg) were added intofreshly distilled 1,2,4-trichlorobenzene (20 ml) and sonicated in anultrasound bath for 10 min. The solution was filtered with filter paper(1#, Whatman) and stored in a glove box (with moisture under 0.5 ppm andoxygen under 0.5 ppm). The working solutions were prepared by dilutingthe stock solutions with TCB.

1.4 Concentration of the Stock Solutions

Since the UV absorbance of TCB overlaps that of nucleosides,concentrations of the stock solutions were determined through a solventexchange. TCB was removed from an aliquot of the stock solution (1 ml)under vacuum at 80° C. and the residue was re-dissolved in the samevolume of chloroform, and its UV absorbance was measured to determineconcentration.

The UV extinction coefficients of all nucleoside derivatives inchloroform were determined at their maximum absorption wavelengths usinga series of dA, dG, dT, and dC dilutions, respectively. The dilutionfactor varied from 3.5 to 200. The curve fitting was carried out inOrigin 8. The resultant concentrations of the stock solutions are listedin the following table.

TABLE 1 Saturation concentrations of nucleosides in TCB stock solutions.dA dG dT dC Wavelength (nm) 260 257 268 280 ε 12690 11470 9480 7400C_(sat) (10⁻⁶ mol/L) 5.8 ± 0.2 2.9 ± 0.1 7.7 ± 0. 1 60.4 ± 7.8 C (10⁻⁶mol/L)  0.7 ± 0.02 2.9 ± 0.1  4.3 ± 0.06  0.8 ± 0.1 The bottom row liststhe final concentrations used in the tunneling measurements(concentrations that resulted in approximately equal read rates for eachnucleoside).

Example 2 Preparation and Characterization of Probes and Surfaces

Gold (S. Chang et al, Nanotechnology 20, 075102 (2009)) (Alfa Aesar,0.25 mm diameter, 99.999% pure) and Pt (20% Ir) (L. A. Nagahara, T.Thundat, S. M. Lindsay, Rev. Sci. Instrum. 60, 3128 (1989)) probes wereetched and the surfaces were prepared (J. A. DeRose, T. Thundat, L. A.Nagahara, S. M. Lindsay, Surf. Sci. 256 102 (1991)) and annealed in ahydrogen flame.

0.3 mg benzoic acid was dissolved in 2 mL N,N-Dimethylformamide(Sigma-Aldrich, >99.99% pure) that was degassed with argon. Substrateswere immersed in this solution for two hours immediately after hydrogenflame annealing, then rinsed with N,N-Dimethylformamide, acetone and1,2,4-trichlrobenzene and dried in flowing N₂ before use.

Prior to modification, probes were cleaned in piranha (H₂SO₄/30% H₂O₂,3:1: creates heat and oxygen—treat with caution). They were thenimmersed into the 1 mM benzoic acid solution overnight, cleaned byN,N-Dimethylformamide, acetone, 1,2,4-trichlrobenzene and blown drybefore use. All measurements were carried out in freshly prepared puresolvent or solutions of nucleosides.

Surfaces were characterized by ellipsometry, STM (A. H. Schäfer, C.Seidel, L. Chi, H. Fuchs, Adv. Mat. 10, 839 (1998)) and FTIR (S. E.Creager, C. M. Steiger, Langmuir 11, 1852 (1995)). The FTIR spectraclearly show that the benzoic acid moiety is exposed and in its neutralform. Background tunneling signals were measured and shown in FIG. 2.

2.1 Polarization Currents and Electrochemical Leakage

The absolute values of peak current are affected by electrochemicalleakage as follows: The tunneling current is set after backing out thebackground leakage current measured with the probe far from the surface.If this is substantial (tens to hundreds of pA) then, even though theprobes are un-insulated (so that leakage is generated over their entiresurface) the leakage can still change (at the pA level) when the probeis brought to the surface as a result of altered diffusion rates aroundthe apex of the probe. Thus the correction applied for leakage with theprobe far from the surface can overcorrect for leakage with the probeclose to the surface. In consequence, the apparent tunnel current isoverestimated, changing the real set-point from nucleoside to nucleosideif the leakage is different from one nucleoside solution to another.This effect is large enough to change the apparent order of the dC anddG peaks at the saturation concentrations (Table 1). Diluted to theworking concentrations shown in Table 1, leakage currents at 0.5V biaswere 1.0 to 2 pA (dA), 0.0 to1.0 pA (dT) and 0.3 to 1 pA (dG). In thecase of dC (0.8 μM) a current of 15 pA was observed initially, but thisfell to a few pA after an hour of exposure to the solution. Thesebackgrounds have been subtracted from the baseline tunnel currentsreported here. They do not appear to cause significant errors asevidenced by the similarity between the data for single nucleosides andthe data for mixtures. Examples of raw data for dT, dG and dC can befound in FIG. 15.

2.2 STM Servo Gain

The frequency response of the servo was determined by comparing 1/fnoise plots without (FIG. 16A) and with (FIG. 16B) the servo applied.

Current traces were Fourier transformed and displayed as a spectraldensity according to:

$\begin{matrix}{{PSD} = {\frac{2\left( {{Re}^{2} + {Im}^{2}} \right)}{n} = {\frac{2}{N\; \Delta \; t}\frac{{Re}^{2} + {lm}^{2}}{f_{n}}}}} & \left( {{Formula}\mspace{14mu} I} \right)\end{matrix}$

where n is the frequency channel number (Δt=20 μs and N=50000). Thesolid line in FIG. 16A is a fit to 1/f. With the servo loop closed (FIG.16B) the noise data is suppressed below 35 Hz, corresponding to a 28 msresponse time. This is long enough not to distort all but the longestpulses (the long pulses in the insets in FIG. 15 show a small fall-offin peak current level consistent with the measured servo response).

2.3 Automatic Peak Detection

Data analysis was automated both for speed and to eliminate operatorbias. The one operator input to the process was to move the probe toquieter areas of the substrate if extremely noisy backgrounds(characteristic of contamination) were encountered.

The principle challenge lay with low frequency instabilities inbackground current that were not completely corrected by the servo. If asmall fixed threshold for acceptance of a peak was used, even a veryshort fluctuation of the baseline above this threshold produced a largenumber of spurious counts. This problem was overcome as follows.

The current-time data (acquired at 50 kHz) were broken into 0.3 sblocks. The amplitudes in a block were binned and the bottom half of thedata fitted by a Gaussian, the program checking that the mean of theGaussian was equal to the desired baseline current. The HWHH of theGaussian was used to determine the SD, σ of the baseline noise. The datashown here were analyzed by setting the threshold to 2σ above the noisein a 0.3 s run of data. Because the noise level varies over the durationof a run, this variable threshold results in a variable cut-off, andwhen data are aggregated, this can alter the shape of the distributionfor the lowest current reads (i.e., for dT and data acquired with oneelectrode bare). The effect of three choices of cut-off on a 30 s run(i.e., 100 0.3 s segments)of data (dT, 4.3 μM, G_(bl)=12 pS V=0.5V) areshown in FIG. 17.

Very short pulses (at the limit of instrumental resolution) can dominatethe data and do not appear to be sensitive to the identity of thenucleoside. Therefore, all spikes of only one (20 μs) or two data pointsduration (40 μs) were rejected. Distributions of spike lifetimes areprovided in FIGS. 27-29.

2.4 Data Obtained with Bare Electrodes

Data obtained with bare electrodes is shown in FIGS. 18 and 19. FIGS. 18and 19 show that distributions are asymmetric. We assume that themolecular configurations are randomly distributed, and that the tunnelcurrent is exponentially sensitive to changes in position. Thus, we useda Gaussian distribution in the logarithm of the current:

$\begin{matrix}{{N(i)} = {N_{0}{{\exp\left( {- \frac{\left\lbrack {{\ln (i)} - {\ln \left( i_{0} \right)}} \right\rbrack^{2}}{\left\lbrack {\ln (w)} \right\rbrack^{2}}} \right)}.}}} & \left( {{Formula}\mspace{14mu} {II}} \right)\end{matrix}$

The equation shown in formula II fits the data well as shown by thecurved lines in FIGS. 18 and 19. The quality of the fit is demonstratedwith a log-log plot of the data from FIG. 19A, shown in FIG. 20. Thecurved line is a parabola.

TABLE 2 Peak currents (I_(p)), widths on the high current side of thedistribution (I_(0.5) ⁺) and read-rates (RR) for the four nucleosidespassing bare gold electrodes set at 20 pS and 40 pS conductance (bias =0.5 V). dT dC dA dG I_(P) I_(0.5) ⁺ RR I_(P) I_(0.5) ⁺ RR I_(P) I_(0.5)⁺ RR I_(P) I_(0.5) ⁺ RR G_(bl) pA pA (s⁻¹) pA pA (s⁻¹) pA pA (s⁻¹) pA pA(s⁻¹) 20 pS 12.4 15.3 0.4 10.0 6.1 0.6 15.9 14.6 5.4 18.7 15.2 4.0 40 pS32.4 22.6 9.2 35.1 21.6 8.8 36.6 70.0 30.6 44.1 37.5 14.7

The differences between Gaussian and Gaussian log fits was less markedfor the narrower distributions measured with functionalized probes,though the fits with the Gaussian log function were still noticeablybetter than fits with Gaussians. Most data were fitted with a sum of twoGaussian, the second being centered at twice the current peak of thefirst:

$\begin{matrix}{{N(i)} = {{N_{1}{\exp\left( {- \frac{\left\lbrack {{\ln (i)} - {\ln \left( i_{01} \right)}} \right\rbrack^{2}}{\left\lbrack {\ln (w)} \right\rbrack^{2}}} \right)}} + {N_{2}{\exp\left( {- \frac{\left\lbrack {{\ln (i)} - {\ln \left( {2i_{0}} \right)}} \right\rbrack^{2}}{\left\lbrack {\ln (w)} \right\rbrack^{2}}} \right)}}}} & \left( {{Formula}\mspace{14mu} {III}} \right)\end{matrix}$

For the narrower distributions, the HWHH is given approximately by

$\begin{matrix}{{\Delta \; i_{\frac{1}{2}}} = {i_{0}\left( {1 - {\exp \left\lbrack {0.8326{\ln (w)}} \right\rbrack}} \right)}} & \left( {{Formula}\mspace{14mu} {IV}} \right)\end{matrix}$

2.5 Data for Mixed Solutions of Nucleosides

Data for mixed solutions of nucleosides is provided in FIG. 21. Readswith mixed films were somewhat heterogeneous, indicative of phaseseparation on the surface. The distributions shown in FIG. 3F and FIG.3H, FIG. 5C and FIG. 21 were obtained by sampling the surface at sixdifferent points and adding the data. For a bulk dA:dG concentrationratio of 0.24 (Table 1) the ratio of measured peak areas is 0.6,suggesting that dA binds the surface with greater affinity than dG. Whenthe bulk concentration ratio is changed to dA/dG=0.12, the ratio of thepeak areas falls only to 0.4, indicative of a complex adsorptionisotherm for the mixture.

For a bulk dT:dC concentration ratio of 0.19 (Table 1) the ratio of theareas of the respective peaks indicates a surface concentration ratio1.1, suggesting that dC has a much higher affinity for this surface.When the concentration ratio is changed to 0.09:1 (FIG. 21) theintegrated peak areas indicate that the surface concentration ratio haschanged to dT/dC=0.2.

Thus in the case of the dC/dT mixed layers, much larger changes inrelative surface concentration result from a given change in the bulkconcentrations than is the case for the dA/dG mixtures, presumably aconsequence of competition between different solvent affinities,different surface affinities and interactions between the nucleosides onthe surface. Nonetheless, the peak associated with the diminishedcomponent is consistently lowered, validating our peak assignments basedon current measurements on pure nucleoside solutions.

2.6 Calculations of Conductance for Hydrogen-Bonded Complexes

The evaluation of the current due to the applied bias is determinedusing ballistic transport theory. The electronic states of the gold leakout across the molecules producing a tunneling current. In-elasticscattering of the electrons during their transmission is not considered.The electronic current is determined by the transmission functionthrough the molecules by electrons at the Fermi level of the metal. Onlyvery small biases are considered (+/−0.1 V). In this region the I-Vcharacteristics were all linear, so that the results are characterizedsimply by its conductivity. The conductivity amounts to the product ofthe quantum of conductance and the transmission function at the Fermilevel.

The computation of the transmission function is given by standardresults from scattering theory (J. K. Tomfohr, O. F. Sankey, J. Chem.Phys. 120, 1542 (2004)), Γ(E)=Tr(Γ_(L)(E)G_(M)(E)Γ_(R)(E)

(E)), where E is the energy (Fermi level of the gold contacts), Γ arethe spectral density of states of the left and right metals contacts,and G_(M) is the molecular Green's function propagator. The Γ functionscontain all the information the metallic states and how they couple tothe molecules and G_(M) contains all the information on the electronicstates within the molecules. The Green's function propagator will decayapproximately exponentially with distance along the path from metalcontact to metal contact.

In order to compute the spectral density of states and Green'sfunctions, one needs a model of the electronic states and methods tomodel the semi-infinite metallic leads. We modeled both the tip and thesubstrate and semi-infinite flat planar gold (111) surfaces.

The connecting sulfur atoms at the termini of the molecules are above Auhollow sites. A supercell slab geometry is used. This means the systemis a periodic array of Au slabs (initially thin) with moleculessandwiched between them in a specific configuration. The repetitivesupercell structure is so that Bloch's theorem can be used to determinethe electronic states of the entire system. The electronic structure ofthe entire supercell is determined self-consistently within densityfunctional theory. To correct for the fact that the slabs are 5-7 layersof Au, a recursion method extends them to infinity by choosing thecentral layer to represent bulk gold.

The electronic structure is determined using local atomic orbitals ofthe fireball (O. F. Sankey, D. J. Niklewski, Phys. Rev. B40, 3979(1989)) type. The local orbitals have a finite radius and are thus veryslightly excited from the ground state. The SIESTA code (P. Ordejon, E.Artacho, J. M. Soler, Phys. Rev. B 53, 10441 (1996)) is used withindensity functional theory. All atoms are described with pseudopotentialswhich eliminates all the core states. The basis set used is double zetaplus polarization (DZP) for all atoms except Au which used a single zetaplus polarization (SZP).

Many different geometries of binding between the two readers and thetarget base were explored. In all cases the relaxed DFT geometry of thereader and base was used. A restricted set of calculations relaxed theentire molecular system. The results reported in Table 3 used geometriesof relaxed individual molecules, and the individual molecules were thenrigidly translated into the assembled structures in FIG. 1A-D. Theimportant variables were the length of the hydrogen bonds (which wereset to expected values for such hydrogen bonded molecules e.g. fromexperiment and DFT (M. H. Lee, O. F. Sankey, Phys. Rev. E 79, 051911 1(2009)), and the distance between the metal leads. The estimatedmetallic lead distance was set to that value estimated from thetunneling decay constant and the background tunneling current though thesolvent.

The quantitative disagreement between theory and experiment (Table 3)appears large—especially so for the case of dT. However, neglect ofsolvent-mediated tunneling probably ignores an important additionalcurrent, equivalent to that detected with just one functionalizedelectrode where the top contact is solvent mediated. This is asignificant current that should be added as a background to thethrough-bond values calculated here. A second source of error probablyoriginates in our estimate of the tunnel gap. A small overestimate (0.1nm out of 2.5) would lead to a significant lowering of the calculatedtunnel current, because of the very large electronic decay constant forstretched hydrogen bonds. M. H. Lee, O. F. Sankey, Phys. Rev. E 79,051911 1 (2009).

Example 3 Tunneling Measurements

We carried out tunneling measurements on a PicoSPM scanning probemicroscope (Agilent, Chandler) interfaced to a digital oscilloscope.When both the probe and a gold (111) substrate were functionalized with4-mercaptobenzoic acid, the tunneling background signal in TCB wasrelatively noise free for set-points currents, I_(bl) of up to 10 pA at0.5V bias, a conductance of 20 pS (FIG. 2). A nucleoside solution wasplaced in the liquid cell, and after the polarization current had fallento a small value we re-engaged the probe at a tunnel current level thathad previously given a low-noise background signal. Current spikes wereimmediately obvious in the tunneling signal (FIG. 15). Because neitherthe surface concentrations of nucleosides nor the efficiency ofmolecular capture in the gap are known a priori, we adjusted theconcentrations of the nucleoside solutions to give approximately equal“spike rates” in the tunnel gap (Table 3).

TABLE 3 Measured and calculated conductances in a functionalized tunneljunction at I_(bl) = 6 pA, V = 0.5 V. dT dG dC dA Measured G 13.6 ± 0.318.6 ± 0.9 25.3 ± 2.5  33 ± 1.9 (PS) Calculated G 0.04 0.12 0.51 1.05(PS) Read rate (s⁻¹)  7.1 ± 1.4  5.5 ± 1.1  5.5 ± 1.1 6.6 ± 1.3 Measuredvalues are the average of three independent runs (errors are ±1 sd).Calculated conductances are for the structures shown in FIG. 1 A-D. Readrate is based on counts acquired in a 180 s period for nucleosideconcentrations between 0.8 and 4.3 μM. The disparity in the range ofvalues between theory and experiment may reflect neglect of a backgroundcontribution via solvent-mediated tunneling into a molecule bound at oneelectrode. Absolute values will be affected by inaccuracies in theestimate of the gap size.

Many of the “spikes” showed the two-level “telegraph noise”characteristic (S. Chang et al., Nanotechnology 20, 075102 (2009)) ofbinding and unbinding of a single molecule in the gap (insets, FIG. 15).The STM servo gains were set so that only spikes of the longest durationwere affected by the action of the current-control servo (FIG. 16).

We generated distributions of the peak currents using a custom programto analyze the height of the spikes. The program captures signals twostandard deviations above the noise on the baseline, and also rejectsdata of only one or two points in time (i.e. up to 40 μs duration). Theeffect of the choice of filtering parameters on the measureddistribution is shown in FIG. 17 and FIGS. 27-28). FIG. 3 shows howthese measured distributions are affected by functionalization of theelectrodes. Distributions recorded with bare electrodes are shown inFIG. 3A and FIG. 3C. In order to record signals with bare electrodes, wehad to reduce the tunneling gap a little by operating at a conductanceof 20 pS. Even at this smaller gap, reads with bare electrodes on thepyrimidine nucleosides were much less frequent than reads on purinenucleosides (FIG. 18, Table 2). The measured current distributions werefitted quite well by a Gaussian distribution of the logarithm of thecurrents (solid lines) as shown in FIGS. 18-20. The fitted peak currentsdiffer for these two nucleosides (15.9±0.4 pA for dA and 18.7±0.2 pA fordG) but the difference (2.8 pA) is less than the width of thedistribution on the high current side (˜15 pA). When measurements arerepeated with a functionalized substrate and a bare gold probe at anincreased gap (corresponding to 12 pS) the distribution of measuredcurrents narrows by an order of magnitude (FIG. 3B—dA), (FIG. 3D—dG) butthe peak currents are not significantly different. The distribution ofspike lifetimes is quite similar for both bare electrodes and for onefunctionalized electrode (FIG. 29). Thus, it appears likely that thespikes observed with bare electrodes correspond to transiently boundstates of the nucleosides also. If this is the case, then the narrowingobserved with a functionalized electrode must be a consequence of areduction in the number of types of bound states in the tunnel gap. Whenboth probe and substrate are functionalized, (FIG. 3E—dA, FIG. 3G—dG),the peak current for dA is clearly higher than the peak current for dG.Thus distinctive signals can be generated when both electrodes arefunctionalized, but do they originate with single nucleosides? The“telegraph noise” signals are characteristic of single-molecule readsand the small size of the peaks assigned to two-molecule reads (“2” inFIG. 3B, FIG. 3D, FIG. 3E, FIG. 3G) suggests that reads of more than onemolecule at a time are infrequent. However, electrochemical leakagecurrents can introduce current errors that depend on the nucleoside sothe measured current may not be generated from single molecule currentsalone. A better test of the fidelity of tunneling reads can be carriedout using mixtures of two nucleosides so that any errors owing to anelectrochemical background are present in both sets of signals. FIG. 3Fshows the current distribution obtained with a mix of dA and dG. Thehigher current peak is at essentially the same current as recorded fordA alone, and thus should count the dA molecules in this mixture. Thisassignment is confirmed by halving the concentration of dA in solution(FIG. 3H). Surface concentrations are unknown a priori and dependent oncompetition between nucleosides for surface binding sites and thedifferent dissociation rates back into solution so absolute signal ratescannot be interpreted in terms of concentrations quantitatively. Most ofthe data in this panel were well fitted assuming single molecule readswith only 5% of the reads consistent with both dA and dG in the gap atthe same time (“dA+dG”, FIG. 3F).

The same types of features are observed for dC and dT (FIG. 5 and FIG.29) but the data for a bare gap and bare substrate had to be collectedat a yet larger tunnel current (20 pA, corresponding to 40 pS) in orderto acquire a significant number of reads for the (smaller-sized)pyrimidine nucleosides (FIG. 19). dC and dT are also clearly separatedin a mixed sample when read with probes that are functionalized (FIG. 5Cand FIG. 21).

At a given bias, the absolute value of peak current is directlyproportional to the baseline conductance of the gap (FIG. 7A), i.e., itincreases exponentially as the gap is decreased, similar to what hasbeen reported for other hydrogen-bonded systems in large tunnel gaps (S.Chang et al., Nanotechnology 20, 075102 (2009)). We found evidence of aninteresting dependence of the peak currents on bias at a fixed gap size(i.e., gap conductance) (FIG. 22) indicating the possibility of anon-linear current-voltage dependence for molecules bound to bothelectrodes. The read frequency also increased as the gap was narrowed(FIG. 24). On the other hand, the fraction of multi-molecule readsincreased rapidly in smaller gaps (FIG. 7A and FIG. 25) so 12 pS appearsto be an optimal value for the baseline conductance at a bias of 0.5V.

Values for the peak currents measured at I_(bl)=6 pA, V=0.5V aresummarized by the cross-hatched bars in FIG. 7B. These are the resultsof three different runs (one carried out, from sample preparation todata analysis, by a different team) on each of the four nucleosides. Thepeaks for each nucleoside are separated by an amount comparable to thewidth of the distribution, allowing the fraction that aresingle-molecule reads with two “good” contacts to identify the base withp≧0.6 (FIG. 26).

We also recorded data with a functionalized substrate and a bare Au(dark shaded bars) or bare Pt (light shaded bars) probe. The peakcurrents change little from nucleoside to nucleoside, an expectedconsequence of the resistance, R_(c), associated with bare contacts (X.D. Cui et al., Science 294, 571 (2001)) although the lack of selectivityis not accounted for by contact resistance alone. If we assume thatreads with two functionalized probes determine a resistance for a singlemolecule, R_(m), then the resistance of a junction with one bareelectrode should be given by R_(j)=R_(c)+R_(m). FIG. 7B shows that thesignal with one bare gold electrode is insensitive to the molecularresistance, while a bare Pt electrode is about half as sensitive as thesimple “resistors in series” model predicts, probably reflecting the wayin which binding to the electrodes affects the position of molecularstates (V. Meunier, P. S. Krstić, J. Chem. Phys. 128, 041103 (2008).

At 12 pS conductance, we estimate the gap to be about 2.5 nm, usingG=G₀exp(−βx) where G₀ is the quantum of conductance (77 μS) and β=6.4nm⁻¹ (J. He, L. Lin, P. Zhang, S. M. Lindsay, Nano Letters 7, 3854(2007)). FIGS. 1A-D show what we believe to be the most likely hydrogenbonded (energy-minimized) structures for the four nucleosides in a gapwith both electrodes functionalized. We carried out density functionalcalculations of the conductance of these four molecular junctions andthe predicted conductances are listed below the measured values in Table3. The predicted order of conductance agrees with experiment, though theabsolute values are significantly lower, possibly because of anoverestimate of the size of the tunnel gap. Lifetime data (FIG. 29) fordT show little change when both electrodes are functionalized, so the dTspikes may represent solvent-mediated tunneling at one electrode. Sincesolvent molcules are not included in the simulations, this additionaltunneling contribution is absent from the predictions. Accordingly, aconstant background should be added to each of the predicted currents,which would diminish the discrepancy between the range of predicted andmeasured currents.

Example 4 Synthesis of 4-Mercaptobenzamide

The synthesis of 4-mercaptobenzamide was carried out according to thefollowing scheme:

4.1 Materials and Methods

Proton NMR (1H) spectra were recorded at 400 MHz on a Varian 400 MHzspectrometer or at 500 MHz on a Varian 500 MHz spectrometer, and carbonNMR (13C) spectra were recorded at either 100 MHz on a Varian 400 MHzspectrometer or at 125 MHz on a Varian 500 MHz spectrometer. HRMSspectra were acquired using the atmospheric pressure chemical ionization(APCI) technique. Flash chromatography was performed in CombiFlash Rf(Teledyne Isco, Inc.). All reagents were purchased from Aldrich unlessotherwise stated.

4.2 Step 1: 4-Tritylmercaptobenzoic Acid

4-Mercaptobenzoic acid (1.54 g, 10 mmol) and trityl chloride (2.79 g, 10mmol) were dissolved in DMF (25 mL) and stirred at an ambienttemperature for 36 h. The solvent was removed under reduced pressure.The residue was dissolved in chloroform (50 mL), and washed with water(3×25 mL). The organic layer was dried over MgSO4, filtered, andconcentrated. Compound 1 was obtained as a white solid (3.20 g, 81%). 1HNMR (500 MHz, CDCl3): 7.67 (d, 2H), 7.21-7.39 (m, 15H), 6.99 (d, 2H).

4.3 Step 2: 4-Tritylmercaptobenzamide

Ammonia (0.5 M in dioxane, 1 mmol) was added drop-wise to a solution ofcompound 1 (198 mg, 0.5 mmol), 1-hydroxy-benzotriazole (HOBt) (68 mg,0.5 mmol), and 1,3-dicyclohexylcarbodiimide (DCC) (103 mg, 0.5 mmol) inTHF (5 mL) at 0° C. The resulting mixture was allowed to warm to roomtemperature, stirred for 24 h. After filtration, the filtrate was washedwith saturated aqueous NaHCO3. The organic layer was dried over MgSO4,filtered, and concentrated. The residue was purified by flashchromatography (silica gel, Dichloromethane: Methanol gradient 100:0 to100:3) to yield compound 2 as a white solid (154 mg, 78%). 1H NMR (400MHz, CDCl3): 6.98-7.43 (m, 19H), 5.95 (brs, 1H), 5.75 (br s, 1H); HRMS(APCI+): found, 396.1442; calcd for C26H22NOS+H, 396.1422.

4.4 Step 3: 4-Mercaptobenzamide

Compound 2 (60 mg, 0.15 mmol) was dissolved in a mix of trifluoroaceticacid (TFA) (2 mL) and triethylsilane (TES) (2 mL), stirred for 2 h atroom temperature. The solution was rotarily-evaporated to dryness underreduced pressure. The residue was crystallized from the mixture ofhexanes and dichloromethane (v:v=1:1) to yield compound 3 as a whitesolid (12 mg, 52%). 1H NMR (500 MHz, CDCl3): 7.68 (d, 2H), 7.31 (d, 2H),6.50 (br s, 1H), 6.29 (br s, 1H), 3.61 (s, 1H); 13C NMR (125 MHz,CDCl3): 169.9, 138.0, 129.4, 128.5, 128.2.4 HRMS (APCI+): found,154.0326; calcd for C7H7NOS+H, 154.0326.

Example 5 Production, Functionalization, and Characterization ofElectrodes 5.1 Electrode Production

Gold tips were electrochemically etched from gold wires (Aesar 99.999%pure) using a mix of HCl and Ethanol (volume ratio 1:1). Only sharp tips(judged by optical microscopy with 300× magnification) were selected forthe insulating process. High Density Polyethylene (HDPE) was used asinsulation. Prior to insulation, gold tips were cleaned with piranha(mixture of oxygen peroxide and sulfuric acid, volume ratio of 1 to3—Caution—this material can explode in reactions with organic materials)for 1 min to get rid of organic contaminations and rinsed with doubledistilled water, ethanol and blown dry with compressed nitrogen gas.During insulation, the HDPE was melted at 250° C. on a homemade tipcoating instrument. Penetration through the melted HDPE will cover mostarea of the tip with the insulating material, leaving only the apexun-insulated. The exposed surface area of insulated tips wascharacterized by cyclic voltammetry in potassium ferricyanide. Insulatedand un-insulated tips were tested for cyclic voltammetry. The insulatedtip provided for a more consistent and regular electrode. FIGS. 30 and31 illustrate these results. FIG. 30 shows cyclic voltammetry for a baregold wire in 50 mM potassium ferricyanide (potential vs. Ag wire). FIG.31 shows cyclic Voltammetry for a HDPE coated STM tip. Assuming ahemispherical exposed tip shape and using the formula Imax=2 πRnFCD, thetypical exposed surface area of the coated scanning probes is on theorder of 10-2 μm.

5.2 Functionalization

Gold substrates were annealed in a hydrogen flame to get rid ofcontamination and form well ordered Au surfaces. 4-Mercaptobenzamideprepared according to Example 4 was dissolved (1 mM) in methanol anddegassed using argon to avoid oxidation of thiols. The insulated tipstreated substrates were immersed in this solution for >2 hours. Thisresulted in the formation of monolayers of benzamide on the surface.Extended functionalization times degraded insulation on the probes sotreatment of probes was limited to 2 h. Functionalization of goldsubstrates was carried out for up to 20 hours.

The thickness of the molecule SAM after deposition was measured byellipsometry (Gaertner, Skokie, Ill.) at a wavelength of 632.8 nm withan incident angle of 70 degrees. The optical constants of the freshlyhydrogen-flamed bare gold substrate (200 nm thick on mica) were measuredbefore deposition of molecules given n=0.2 and k=−3.53. The SAM opticalconstants were set to nf=1.45 and kf=0. The thickness of4-mercaptobenzamide monolayer was measured as 0.70±0.17 nm.

Infrared absorption spectra of the SAM were recorded using the SmartApertured Grazing Angle accessory on Thermo Nicolet 6700 FTIR (ThermoFisher Scientific, Mass.). The spectrum of the powder sample was takenusing Smart Orbit (a diamond single-bounce ATR accessory). FIG. 32 showsFTIR spectra of the 4-mercaptobenzamide monolayer (lower line) andpowder (upper line). In the monolayer IR, absorption peaks at 3487 cm-1,1610 cm-1, and 1404 cm-1 are assigned as N—H stretching of discrete NH2(without hydrogen bonds), amide band I, and amide band II, respectively.

The ellipsometry data suggested almost a full monolayer coverage,however, STM images showed (FIG. 33) that the coverage occurs inpatches. The read rates recorded in the paper are for reads taken withthe probe positioned over functionalized patches. In FIG. 33, the STMimage shows islands of mercaptobenzamide on Au(111) surface. (Image in 1mM PB buffer with a gold tip, 0.5 volts tip bias with 10 pA set point.)

5.3 Imaging the Electrodes

The electrode was characterized by optical microscope and Transmissionelectron microscope (TEM) image. FIGS. 34 a-c show a bare electrode andFIGS. 34 d-f show a polyethylene coated tips. FIG. 34 a shows an opticalmicroscope image of a typical “good” tip. This tip was furthercharacterized by transmission electron microscopy (TEM) as shown inFIGS. 34 b-c. The tip radius in this case is about 16 nm (FIG. 34 c).The carbon layer (the white layer covering the gold tip) was depositedduring TEM imaging. The dashed arc has a radius of 16 nm. The measuredradii typically spanned the range from 5 to 20 nm. The tip surface isnormally smooth, but bumps (1-2nm in height) are observed sometimes. Itis, at first sight, surprising that such “blunt” probes could yieldsingle molecule resolution, but the adsorption of molecules onto thesurface generates local high points capable of single moleculeresolution (and even better as illustrated by the resolution of internalmolecular structure with functionalized AFM probes).

An optical microscope image of a typical insulated tip is shown in FIG.34 d. This tip showed no leakage current (below the measurement limit ˜1pA) and about 8 pA (peak to peak, i.e. 4 pA above the baseline) 120 Hznoise in experiments. The TEM images of the same tip are shown in FIGS.34 d-f. The arrows in FIGS. 34 e-f indicate the exposed gold (highresolution imaging is not possible owing to charging of the coating).

Example 6 Characterization of the Tunnel Gap 6.1 Water and Buffer

A tunnel gap with electrodes as described in Example 5 was characterizedusing doubly distilled water and 0.1 mM phosphate butter (PB—pH=7.4).Small signals were observed from the buffer alone with bare electrodes,but they were much rarer when both electrodes were functionalized andthe tunnel gap conductance set to 20 pS or less. See FIG. 55. The tunneldecay was much more rapid (decay constant, β=14.2±3.2 nm⁻¹) with bothelectrodes functionalized than is the case in water alone (β˜6.1±0.7nm⁻¹-¹¹). See Example 6.2 below. It is estimated the tunnel gap at i=10pA and V=+0.5V is a little over the length of two benzamide molecules(i.e. a little greater than 2 nm).

6.2 Background Water Signals

With a gap size of 20 pS at 0.5 volts, control experiment with bareelectrodes on a functionalized substrate in doubly distilled water givebackground telegraph noise signals of a small amplitude (around 6 pA at0.5 volts bias—FIG. 35). However, with functionalized tips onfunctionalized substrate, such signals are generally not observed(occasional observations of signals may originate with incompletecoverage of 4-mercatobenzamide on the surface of either the tip or thesubstrate). These background signals can be excluded by a thresholdvalue in the data analysis since they are smaller in magnitude and lessfrequent than the DNA signals.

6.3 Tunneling Decay Curves

Decay curves were measured in doubly-distilled water with a combinationof functionalized and non-functionalized electrodes. The decay constant(β) was calculated from the slope of a linear fit of a plot of Ln (I)vs. distance (FIG. 36). FIG. 36 shows tunnel current decay curves inpure H2O (multiple curves are plotted in each case). FIG. 36 a showsbare gold electrodes. FIG. 36 b shows both electrodes functionalized.FIG. 36 c shows one electrode bare and the other functionalized. Asignificant increase in β is detected for functionalized electrodes atlarge distances indicating a change of gap composition that we take tobe a transition from the region where the benzamides interact to onewhere they do not. The distributions of measured vales of β are shown inFIG. 37. FIG. 37 shows histograms of beta in pure water for (a) baregold electrodes; (b) both electrodes functionalized withmercaptobenzamide; and (c) one electrode functionalized and the otherbare. Gaussian fits (mean±SD) yield: (a) 6.11±0.68 nm⁻¹ (b) 14.16±3.20nm⁻¹ (c) 6.84±0.92 nm⁻¹.

Example 7 Analyzing DNA Nucleotides in a Tunnel Gap

DNA nucleotides (10 μM in PB) were introduced into a tunnel gap createdusing electrodes as described in Example 5 in an aqueous electrolytesolution. These nucleotides yielded characteristic noise spikes as shownin FIGS. 56 c-f. The signal count rate (defined in FIG. 57) variedconsiderably from 25 counts/s (5-methyl-deoxycytidine 5′-monophophate,dmCMP) to less than 1 c/s (deoxycytidine 5′-monophophate, dCMP). Nosignals were recorded at all with thymidine 5′-monophophate (dTMP), thesignal looking exactly like the control (FIG. 55). STM images suggestthat this nucleotide binds to the surface (and presumably the probe)very strongly, blocking interactions in which a single molecule spansthe junction.

The current occurs in bursts of spikes (longer signal runs are shownhereafter) and distributions of the spike heights were quite well fittedwith two Gaussians distributions of the logarithm as shown in FIGS. 56g-j (the fitting parameters are described in Example 8 below). Thesehistograms were generated by counting only pulses that exceeded 1.5× theSD of the local noise background—i.e., typically pulses above 6 pA (afull description of the analysis procedure is given by Chang et el.).

dCMP generates the highest signals and the lowest count rate whiledeoxyadenosine 5′-monophophate (dAMP) and dmCMP produce the smallestsignals and the highest count rate. Little difference was found betweencytidine and 5-methylcytidine in organic solvent as discussed Example 9below. The three bases with narrower pulse height distributions (dAMP,dmCMP and GMP) often show bursts of “telegraph-noise” characteristic ofsources that fluctuate between two levels (particularly marked fordAMP). Such a two-level' distribution is a strong indication that thetunneling signals are generated by a single molecule trapped in thetunnel junction. The characteristics of the tunneling noise from thenucleotides are summarized in Table 4.

TABLE 4 Nucleotide tunneling noise characteristics. Parameters aredefined in FIG. 57. Nucleotide dAMP dGMP dCMP d^(m)CMP Burst Dura- 0.19± 0.05* 0.13 ± 0.02* 0.12 ± 0.02* 0.06 ± 0.01* tion (T_(B) , s) Burst732 ± 82^(§)  574 ± 67^(§)  306 ± 23^(§)  1305 ± 100^(§)  Frequency(f_(B,) Hz) Fraction of 0.02 0.001 0.02 0.01 reads >0.1 nA τ_(on) (ms)0.38 ± 0.01* 0.48 ± 0.02* 0.42 ± 0.02* 0.31 ± 0.09* τ_(off) (ms) 0.35 ±0.01* 0.56 ± 0.04* 0.71 ± 0.06* 0.41 ± 0.11* τ_(on)/τ_(off) ~1 0.9 0.60.8 ΔG 0 0.1 0.51 0.22 (kT units) *Error in fit to exponentialdistribution. ^(§)Standard error

dAMP signals are well-separated from dCMP signals, and dmCMP signals arewell separated from dCMP signals in spike amplitude and in the timedistribution of their signals (Table 4 and discussed hereafter). Forthis reason, DNA oligomers composed of A, C and mC bases were furtherinvestigated below in Example 11.

Example 8 Gaussian Fits to Current Distributions

Peaks generated from the data of Example 7 were fitted with a Gaussiandistribution in the logarithm of the tunnel current, a model thatassumes a random distribution of tunnel geometries is sampledexponentially. For the present data in water, two peaks were required,implying two binding geometries:

$\begin{matrix}{{N(i)} = {{N_{1}{\exp\left( {- \frac{\left\lbrack {{\ln (i)} - {\ln \left( i_{01} \right)}} \right\rbrack^{2}}{\left\lbrack {\ln \left( w_{1} \right)} \right\rbrack^{2}}} \right)}} + {N_{2}{{\exp\left( {- \frac{\left\lbrack {{\ln (i)} - {\ln \left( i_{02} \right)}} \right\rbrack^{2}}{\left\lbrack {\ln \left( w_{2} \right)} \right\rbrack^{2}}} \right)}.}}}} & {{Equation}\mspace{14mu} S\; 1}\end{matrix}$

Fitting parameters are listed in Table S1.

TABLE S1 Intensity Distribution Fitting Parameters Sample i₀₁ w₁ I₀₂ w₂N₂/N₁ damp 0.014 0.712 0.022 0.491 0.45 d(A)₅ 0.013 0.727 0.022 0.5310.43 dCMP 0.032 0.880 0.042 0.709 0.64 d(C)₅ 0.028 0.787 0.044 0.8350.32 d^(m)CMP 0.017 0.771 0.024 0.627 0.62 d(^(m)C)₅ 0.013 0.723 0.0190.443 0.35 dGMP 0.016 0.852 0.022 0.596 2.01

Example 9 Current Distributions for Cytidine and 5methylcytidine inOrganic Solvent

Data for mC measured in organic solvent with electrodes and4-mercaptobenzamide readers according to Examples 4-5 are included hereto show that the overlap between the signals from these two bases ismuch greater in organic solvent, which demonstrates that water moleculesplay a role in generating different signals from C and meC in thepresent work. FIG. 40 shows current distributions measured for cytidine(solid) and ^(5me)cytidine (hashed) using benzoic acid readers intrichlorobenzene solvent.

Example 10 Analyzing Single Base Within a Heteropolymer 10.1 Reading aSingle Base Within a Heteropolymer

A d(CCACC) oligomer was analyzed by a two electrodes, a probe and asubstrate both functionalized with 4-mercaptobenzamide as describedabove in Examples 4-5. Characteristic bursts of current were observed,and an example of which is shown in FIG. 54 b (the spike labeled “*” isnon-specific and rejected from the analysis). The background tunnelcurrent is 10 pA, bias+0.5V. As shown below, the low frequency, largeamplitude pulses indicate a C, while the high frequency, small amplitudepulses signal an A. FIG. 54 c shows a sliding average of the spikeamplitudes (0.25 s window, 0.125 s steps). Values below the straightline identify an A base unambiguously. FIG. 54 d shows a sliding averageover the pulse frequencies (as defined for each adjacent pair ofspikes)—the low frequency regions at each end enhance the confidencewith which those, regions can be assigned to a C base. C bases generatea negligible number of spikes below 0.015 nA (red line). The probabilityof an assignment to A or C is shown in FIG. 54 e. Calculation of theseprobabilities is based on studies of nucleotides, homopolymers andheteropolymers as described herein. This example clearly shows that asingle A base can be identified with high confidence when flanked by Cbases in an intact DNA molecule.

10.2 Current Traces Showing Bursts for Nucleotides and Longtime Tracesfor d(CCACC)

FIG. 38 shows a sample of tunneling noise generated over a 10 s periodas the probe drifts over a surface covered with d(CCACC). FIG. 39 showstypical “bursts” of signal from each of the nucleotides. In FIG. 38, atypical 10 s time trace for d(CCACC) is shown. Note the preponderance ofA signals. The current spike distribution (inset) is almost completelydominated by “A” signals with the C component in the fit (see curvesmaller box) being 7% or less. This shows that the probe spends moretime bound to the minority of A bases. In FIG. 39, there are longer timetraces for the nucleotides showing typical bursts of data. Each of theseexamples is surrounded by spike-free regions of current.

Example 11 Analyzing DNA Oligomers with4-Mercaptobenzamide-Functionalized Electrodes

FIGS. 58 a,c and e show representative tunneling noise traces for d(A)5,d(C)5 and d(mC)5 with the corresponding current peak distributions shownin FIGS. 58 b, d and f. Comparing FIG. 58 b (d(A)5) with FIG. 56 g(dAMP), FIG. 58 d (d(C)5) with FIG. 56 h (dCMP) and FIG. 58 f (d(mC)5)with FIG. 56 i (dmCMP) demonstrates that surprisingly, most of thepolymer binding events in the tunnel junction generate signals thatresemble those generated by single nucleotides. This finding suggeststhat (1) that single bases are being read and (2) that stericconstraints owing to the polymer backbone do not prevent base-bindingevents from dominating the signals.

There are some (small) differences between nucleotide and oligomersignals, these are: (1) Peak positions, widths and relative intensitiesare somewhat altered. (2) Almost all of the signals generated bynucleotides are less than 0.1 nA at 0.5V bias (Table 4). In contrast,20% of the total signals generated by d(A)5 and d(mC)5 are larger than0.1 nA at this bias (Table 5 (discussed below)—this is not obvious inFIGS. 56 G-J where distributions are plotted only up to 0.1 nA). Thesehigh current (>0.1 nA) features in d(A)5 and d(C)5 are continuouslydistributed so they do not represent parallel reads of more than onebase at a time (where currents would be distributed in multiples of thesingle molecule values). Rather, they are new features associated withthe presence of the polymeric structure in the tunnel gap. Such anon-specific, large amplitude spike is labeled by an asterisk in FIG. 54b.

Features at I>0.1 nA appear much less frequently in oligomers of mixedsequence, suggesting that they are associated with base-stacking in thehomopolymers. FIG. 58 h shows a current distribution for d(ACACA) where95% of events are below 0.1 nA. FIG. 58 j shows a current distributionfor d(CmCCmCC) where 99% of events are below 0.1 nA. The solid red linesare the sums of the distributions measured for the homopolymerscorresponding to the constituents with, scaling aside, only one fittingparameter. This parameter is the ratio, rfit, of the A/C (rfit=0.48) ormC/C (rfit=0.66) contributions. These values differ from the knowncomposition ratios (0.6 for ACACA and 0.4 for CmCCmCC) but aresurprising in as much as the spike rate for dCMP alone is very small,yet C appears to be quite well represented in the mixed sequenceoligomer data. This suggests that Cs surrounded by As are read morefrequently, possibly because the C-containing oligomer is betterattached to the substrate than the isolated dCMP.

Importantly, mixed oligomers generate signals that are largely describedas the sum of the individual base signals. (Some intermediate currentreads, labeled “1” in FIGS. 58 h and j, and a small number of additionalhigh current features—labeled “2”—show that sequence context plays asmall role.)

In these experiments, the probe drifts randomly over the samples, so thesequence is not “read” deterministically. Nonetheless traces in whichthe signals alternate between “A-like” and “C-like” (FIG. 58 g) and“mC-like” and “C-like” (FIG. 58 i) may be readily found. The duration ofthese “bursts” (see FIG. 57 and Example 11) of signals is long(0.14±0.02 s in ACACA and 0.15±0.02 s in CmCCmCC). Similar bursts areseen in the homopolymers (Table 5) and the nucleotides (Table 4).

TABLE 5 Oligomer tunneling noise characteristics. Parameters are definedin FIG. 57. Oligomer d(A)₅ d(C)₅ d(^(m)C)₅ Burst Duration 0.14 ± 0.02*0.15 ± 0.03* 0.41 ± 0.03* (T_(B,) s] Burst 738 ± 100^(§) 320 ± 85^(§) 662 ± 116^(§) Frequency (f_(B) Hz) Fraction of 0.20 0.0 0.23 reads >0.1nA τ_(on) (ms) 0.33 ± 0.01* 0.34 ± 0.02* 0.26 ± 0.01* τ_(off) (ms) 0.52± 0.02* 0.42 ± 0.01* 0.47 ± 0.01* τ_(on)/τ_(off) 0.6 0.8 0.6 ΔG (kTunits) 0.51 0.22 0.51 *Error in fit to exponential distribution.^(§)Standard errorThis leads to another unexpected finding, namely that the lifetime ofthe bound complex in the tunnel gap is very long (fraction of a second)compared to either the interval between noise spikes (ms) or thelifetime of the bound-state in solution (very short).

Example 12 Distributions of T_(on) and T_(off) of Example 11

Current spikes with durations of <0.1 ms are distorted by the slow (10kHz) response of the current to voltage converter, while pulses of morethan a few ms duration are affected by the feedback used to maintain thetunnel gap. The distributions of t_(on) are shown for monomers in FIG.41 and oligomers in FIG. 42. Distributions of t_(off) are shown formonomers in FIG. 43 and oligomers in FIG. 44. The solid lines are fitsto an exponential decay:

${N(t)} = {\exp \left( \frac{- t}{\tau} \right)}$

giving the ton and τoff values listed in Tables 4 and 5.

FIG. 41 shows the distribution of on times for dGMP, dCMP, dAMP, anddmCMP. Solid lines are exponential fits (from the top: 1^(st) line isGMP, 2^(nd) line is CMP, 3^(rd) line is AMP, and 4^(th) line is 5 meC).FIG. 42 shows the distribution of on times for d(C)5, d(A)5 and d(mC)5.Solid lines are exponential fits (from the top: 1^(st) line is CCCC,2^(nd) line is AAA, and 3^(rd) line is 5 mCCC). Distributions are lesswell separated than in the monomers. FIG. 43 shows the distribution ofoff times for dGMP, dCMP, dAMP and dmCMP. Solid lines are exponentialfits (from the top: 1^(st)line is CMP, 2^(nd) line is GMP, 3^(rd) lineis 5 meC, and 4^(th) line is AMP). FIG. 44 shows the distribution of offtimes for d(C)₅, d(A)₅ and d(mCMP)₅. Solid lines are exponential fits(from the top: 1^(st)line is AAAAA, 2^(nd) line is 5 mCCCC, and 3^(rd)line is CCC). Again, distributions are less well separated than in themonomers.

Example 13 Testing the Long Lifetime of the Complex

Dynamic force spectroscopy was used as an independent test of theunexpectedly long lifetime of the4-mercaptobenzamide-base-4-mercaptobenzamide complex confined to ananoscale gap prepared according to Examples 4-5. In these measurements(FIG. 59 a) one of the recognition molecules was bound to an AFM probevia a 34 nm long polyethyleneglycol (PEG) linker while the other formeda monolayer on an Au(111) substrate. dAMP was used as the target analyteto bridge the gap. In the absence of dAMP, adhesion between probe andsubstrate was extremely small, presumably because the hydrogen bondingsites on the benzamide recognition molecules were stably bound by watermolecules. Adhesion features were observed in the presence of a smallamount of dAMP, falling as the concentration of dAMP increased(resulting in binding of both probe and substrate by dAMP). Stretchingof the PEG tether generated a characteristic signal that permittedmultiple binding events (FIG. 59 b (i)) to be separated from singlemolecule events (FIG. 59 b (ii)) so that only single-moleculebond-breaking events were analyzed. Single molecule bond-breaking forcesas a function of pulling speed are summarized in FIG. 59 c (solid linesare maximum likelihood fits to a heterogeneous bond model) and the bondsurvival probability as a function of bond-breaking force is shown inFIG. 59 d. The solid lines are fits to the same heterogeneous bondmodel. They yield an off-rate at zero force:

K _(off) ⁰=0.28 s⁻¹.

Thus, the intrinsic (zeroforce) survival time of this complex is on theorder of seconds, not milliseconds. The analysis also yields thedistance to the transition state for dissociation, α=0.78 nm (as well asits variance, σ=0.19 nm). Its thus concluded that each base resides inthe tunnel junction for a significant fraction of a second, whilegenerating tunneling signals at kHz rates. Thus the entire cluster ofsignals that occur in one burst (burst durations are listed in Tables 4and 5) can be used to characterize a base.

Long-bound-state life-times accompanied by rapid fluctuations inelectronic signatures have been reported previously in STM images and inthe effect of single-molecule reactions on transport in carbonnanotubes. The origin of this noise is unclear, save that it appears tobe very temperature sensitive, indicative of small energy barriers tothe motion that causes the noise. The distribution of “on” and “off'times were analyzed (see FIG. 57). In a limited time range of times,determined by the amplifier response at one end, and the servo responsetime at the other, these distributions are exponential (as expected fora Poisson source) and the 1/e times (τon and τoff) are listed in Tables4 and 5. They do not differ much, and calculating an energy difference,ΔG, between the on and off states from:

ΔG=kT _(B) ln(τ_(off)/τ_(on))

yields the values listed in Tables 4 and 5 (in units of thermal energy,k_(B)T at 300K). These values are all a fraction of k_(B)T. Thus the“switching” cannot represent thermal activation over a significantbarrier (the normal source of two-level noise). One possible explanationis Brownian motion in a bound state sampled by anexponentially-sensitive matrix element.

The “on” and “off” times are so broadly distributed that they are notvery useful for identifying base-signals. However, the frequency withina burst (f_(s) Tables 4 and 5) is a much simpler parameter. FIGS. 49 and50 show the current distributions and frequency distributions for thethree homopolymers, normalized so that the area under each curve isunity. The frequency distribution for d(mC)5 is bimodal, with many readsin the “C” frequency range and a number at the very fast rate (ca. 1300Hz) observed for mC MP alone (labeled f(mCMP) on the figure). Thissuggests that the binding modes of mC are altered significantly in apolymer context (consistent with the larger shift of the polymer signalcompared to the nucleotide signal, FIG. 58 f) so we chose to analyzeoligomers containing A and C, in particular the d(CCACC) sequencepreviously analyzed.

Given an average current in a burst,

i

and frequency,

fΔ the distributions shown in FIGS. 52 a-b,

-   -   I_(A,C)(        i        ) (FIG. 52 a) and F_(A,C)(        f        ) (FIG. 52 b)        determine independent probabilities that a base is an A or a C:

$P_{A,C}^{i} = {{\frac{I_{A,C}\left( {\langle i\rangle} \right)}{{I_{A}\left( {\langle i\rangle} \right)} + {I_{C}\left( {\langle i\rangle} \right)}}\mspace{14mu} {and}\mspace{14mu} P_{A,C}^{f}} = {\frac{F_{A,C}\left( {\langle f\rangle} \right)}{{F_{A}\left( {\langle f\rangle} \right)} + {F_{C}\left( {\langle f\rangle} \right)}}.}}$

The current distribution from d(CCACC) (inset in FIG. 38) is almostcompletely dominated by A spikes (the component of the C distribution inthis fit is 7% or less). This is a surprising result, that more C's inthe sequence give a smaller number of C spikes. But it is consistentwith our hypothesis that the frequency of C reads is increased when thebase is flanked by A's (c.f. the increase in C reads in d(ACACA comparedto the dCMP vs. dAMP count rate). Armed with our analysis of the burstsignals, quantitative assignments of mixed signals may be made (this wasdone “by eye” in FIGS. 58 g and i). d(C)5 produces no signals below0.015 nA, so bursts of current below this level (but above the noise)can be unambiguously assigned to A. For larger amplitude signals boththe frequency and amplitude data were used. The result is the pair ofcurves shown in FIG. 54 e. Using this approach to sequence DNA requiresseveral further developments. Firstly, the polymer must be pulledthrough a tunnel junction at a controlled speed, particularly ifhomopolymer runs are to be read. Since DNA passes throughunfunctionalized nanopores too rapidly to be read the long residencetime of bases in a functionalized tunnel junction is an asset. Atpresent, movement from one site to another is driven by uncontrolledmechanical drift that generates unknown forces on the reading complex.Our force spectroscopy data can be used to give a crude estimate of the“pulling” force that would be needed to achieve a given read rate(assuming the measured off-rate for dAMP to be representative for allbases). The Bell equation gives the off rate at a force F as:

$K_{off} = {K_{off}^{0}{\exp \left( \frac{F\; \alpha}{k_{B}T} \right)}}$

with K_(off) ⁰=0.28 s⁻¹ and α=0.78 nm, 19 pN would result in passage of10 bases per second. A rate of 10 bases s⁻¹ gives about 30 data spikes(on average) for a “C” read, enough to generate an assignment with areasonable level of confidence. A force of 19 pN can be generated by abias of just 80 mV across a nanopore 17 so read rates of 10 bases persecond per tunnel junction seem feasible.

Example 14 High Current Tails in the Current Distributions

FIG. 45 shows that distribution of counts for spikes >0.1 nA for d(A)₅for 4-mercaptobenzamide-functionalized electrodes. These are about 20%of the total and are not observed in dNTPs or d(C5). FIG. 46 shows thatdistribution of counts for spikes >0.1 nA for d(mC)5. These are about20% of the total and are not observed in dNTPs or d(C5).

Example 15 SPR Estimation of Interactions of Nucleoside Monophosphateswith 4-mercaptobenzamide on a Gold Substrate and Bound State Lifetime inSolution

Surface Plasmon Resonance (SPR) sensorgrams were recorded on a BI-2000SPR system (Biosensing Instrument, Tempe, Ariz.) that is equipped with atwo-channel flow cell consisting of a polyaryletheretherketone (PEEK)cell block and a polydimethylsiloxane (PDMS) gasket. The wavelength ofthe incident light is 635 nm. Before each experiment, the flow cell wascleaned with ethanol and doubly distilled water.

The SPR sensor chip was fabricated by sequentially coating a 2 nm-thickchromium film and a 47 nm-thick gold film on a BK7 glass cover slide(VWR#48366067) in a sputter coater (Quorum Emitech Corporation, modelK675XD). The gold substrate was cleaned with deionized water, absoluteethanol, nitrogen blowing, and then hydrogen flame annealing before use.A monolayer of benzamide was formed by on line injecting an ethanolicsolution of 1-mercaptobenzamide to the gold chip placed on the SPRinstrument using the serial channel mode. With molecules bonding to thegold surface, the SPR signal increases and eventually reaches a steadyresponse, indicating a maximal coverage of the monolayer. Theinteractions of four naturally occurring nucleoside-5′-monophosphateswith the benzamide surface were measured using a single channel mode onthe SPR instrument. The sample solution injected via an injection valveflowed through one channel, while a PBS buffer (pH 7.4, 10 mM phosphateand 150 mM NaCl) flowed through the other one. The measurements werecarried out at a flow rate of 60 μL min⁻¹ with concentration ofnucleoside monophosphates at 1 mM in the PBS buffer.

The data analysis was carried out in the software provide by the vendor.All data sets were fit to a simple 1:1 interaction model.

The data do not determine K_(off) but the very large values for K_(D)(several mM) imply a rapid off rate. For example, assuming a (small)value of K_(on)=10⁶ M⁻¹s⁻¹, a mM K_(D) yields K_(off)=K_(D)K_(on)=103 orms timescales for the bound state lifetime.

FIG. 47 shows SPR sensorgrams of nucleoside-5′-monophosphates (A, C, G,T, R) interacting with the benzamide surface (R: 2-Deoxyribose5-phosphate sodium salt containing no DNA base). Red lines are fittedcurves modeled to describe a 1:1 binding event. Table 6 shows the rateconstants and dissociation constants derived from the 1:1 bindingkinetic analysis.

TABLE 6 ka kd KD (mM) RSD A 43 (3) 0.159 (2) 3.7 (3) 1.914 C 60 (4)0.181 (3) 3.0 (3) 1.845 G 32 (4) 0.172 (2) 5.3 (6) 1.675 T 68.6 (6) 0.195 (4) 2.9 (3) 2.829

Example 16 Frequency of Bond Breaking Reads in Force Spectroscopy

After testing for interactions in the presence of buffer alone (FIG. 48a—shows the only detected adhesion events out of 1024 pulls), 1 μM dAMPwas added to the liquid cell configured with4-mercaptobenzamide-functionalized electrodes and then force curves weretaken as a function of the number of rinses of substrate and tip with0.1 mM PB. The data show an initial increase as excess dAMP is removed,followed by a decrease with continued rinsing (FIG. 48 b-e).Specifically, FIG. 48 shows (a) control curves taken in the absence ofdAMP showed almost no adhesion events between the benzamide molecules,presumably because they were blocked by water. Addition of dAMP led to anumber of adhesion events that increased as excess dAMP was rinsed outof the system (b,c) decreasing as the rinsing continued (d,e).

Example 17 Noise Model

The purpose of this example is to outline a possible origin of thesignals that are used to identify bases. As shown in example 13, theintrinsic life time of the bound comple in the tunnel junction islong—on the order of seconds. So the base is generally bound for all thetime that it is adjacent to a functionalized electrode if the electrodeor probe is translated such that even a few bases are read per second.What is the origin of the current spikes that repeat on ms timescales?The distributions of “on” and “off” times given in example 12 can beused to compute an energy difference between the “on” and “off ” states.This is listed as ΔG in Table 5. It is a fraction of thermal energy at300K (the units are kT at 300K). Thus the spikes cannot be a consequenceof the molecule jumping between a set of distinct thermally stablestates. Here, we investigate the possibility that continuous Brownianmotion can appear “spiky” when sampled in an exponential way (i.e., bytunneling which is exponentially sensitive to distance). Note that thismodel underlies the choice of an exponential distribution of thelogarithm of currents (equation S1 in example 8). Brownian motion wassimulated with a 1-D,random walker driven by Gaussian (i.e., thermal)noise. The displacement was exponentiated to simulate the effect of atunnel current readout of position. The following MatLab program wasused:

-   -   for x=2:10000    -   z=randn(1);    -   y(x)=correlation*y(x−1)+0.1*z;    -   end    -   a=exp(beta*y);    -   plotyy(t,y,t,a)        The variable “correlation” describes how much of the position on        on one step is retained in the next step. Plots are shown for        various values of the parameter “correlation” in FIGS. 49-51. A        value close to 1 was required to obtain noise spikes that        resemble the observed noise. The Intensity distribution was well        fitted with a Gaussian in the logarithm of current (c.f.,        equation S1) and the time intervals between spikes was        exponentially distributed. FIGS. 49-51 show simulated        displacement (upper reads) and current (lower reads) vs.        time-steps for three values of correlation, C.

Example 18 Probability Calculations

FIG. 52 shows normalized distributions for signals obtained fromhomopolymers in a device with 4-mercaptobenzamide-functionalizedelectrodes. FIG. 52A fits to normalized current distributions (from leftto right: 1^(st) peak=mC, 2^(nd) peak=A, 3^(rd) peak=C). FIG. 52B showsnormalized spike frequencies (fS—see FIG. 57) in a signal burst,measured and fitted with polynomials (line beginning at about 0.2=A,line beginning at about 0.65=C, line beginning at about 0.4=mC). Thefits to the distributions are used to assign the probability that aparticular noise burst originates from an A or a C (if the averagecurrents and frequencies lie above or below the crossover points,labeled “I_(AC)” and “f_(AC)”). Current distributions for C and mC areseparated (crossover=“I_(mC)”) but frequency distributions overlap.

Values of:

-   -   I_(A)(        i        ) and I_(C)(        i        )        are taken from FIG. 52 a. Since there are essentially no current        spikes for C below 0.015 nA, bursts with average intensities        smaller than 0.015 nA can be assigned to A reads. For bursts        with intensities >0.015 nA, we use the values of:    -   F_(A)(        f        ) and F_(C)(        f        )        taken from the normalized distributions FIG. 52 b, calculating        the probability of an A read from the following:

1−P_(C) ^(i)P_(C) ^(f)

and the probability of a C read from the following:

1−P_(A) ^(i)P_(A) ^(f)

Example 19 Synthesis of Imidazole-2-Carboxamide

A short ω-functionalized alkyl is needed to attachimidazole-2-carboxamide to electrodes. Because a variety of4(5)-alkylated imidazoles are reported in literature or are commerciallyavailable, a general method to synthesize imidazole-2-carboxamides byamidation on the imidazole ring was developed. As delineated in thebelow Scheme, 4(5)-(2-thioethyl)imidazole-2-carboxamide (5a) and4(5)-(2-aminoethyl)imidazole-2-carboxamide (5b) were synthesized byamidating 4(5)-(2-(benzylthio)ethyl)imidazole (1a) andN-[2-(4-Imidazolyl]ethyl]phthalimide (1b) respectively. The thiol andamine function as anchor groups for attaching the molecule to metaland/or carbon electrodes. In the same way,4(5)-(tert-butyldimethylsilyloxymethyl)imidazole-2-carboxamide (5c) wassynthesized from 4(5)-(tert-butyldimethylsilyloxymethyl)imidazole (1c),which was used for NMR studies in organic solvents (vide infra).

Two routes to synthesizing these imidazole-2-carboxamides were explored.The 2-position of imidazole can be substituted with formate ester ¹⁵ ora cyano group,¹⁶ both of which can readily be converted into amide. Itwas found that the cyano route gave us the best results. First, compound1a, 1c and 1b were converted into the 1H nitrogen protected products(2a, 2b, 2c) in good yields by reacting with benzyl bromide. NMRconfirms that each of them is a mixture of two isomers. The cyano groupwas introduced into the 2-position of the imidazole ring of 2a, 2b, and2c by treating them with 1-cyano-4-(dimethylamino)pyridinium bromide(CAP). CPA was in situ generated by mixing equivalent amounts ofcyanogens bromide and 4-(dimethylamino)pyridine in diemthyl formamide(DMF) at 0° C. A 2.5 fold of CAP resulted in the best yield. The cyanogroup of 3a, 3b, and 3c was converted into amide (4a, 4b, 4c) in fairyields by hydrolysis in sulfuric acid (20% by volume) andtrifluoroacetic acid (18% by volume). We have tested the basic conditionin the presence of hydrogen peroxide, but it failed to furnish thedesired products. Final products of 5a, 5b, and 5c were obtained byremoving the protecting groups with sodium in liquid ammonia. It isworth noting that the tert-butyldimethylsilyloxymethyl group was stableunder the deprotecting condition. The desired compound 5c was seperatedin a good yield.

Example 20 Analyzing DNA with Imidazole-2-carboxyamide-FunctionalizedElectrodes

Electrodes were prepared and functionalized withImidazole-2-carboxyamide. A fixed tunnel gap was configured anddeoxy-nucleotides were analyzed with baseline tunneling conditions of 6pA, 0.5V. The control group showed almost no signals. FIGS. 62A and Bshow current distributions for ^(m)C, A, T, C, and G. As is apparent,the current signatures for each of the nucleotides are distinct, thusdemonstrating the effectiveness of imidazole-2-carboxyamide as areagent.

Example 21 Analyzing DNA Oligomers withImidazole-2-carboxyamide-Functionalized Electrodes

The electrodes of Example 20 were used to analyze the several homo- andhetero-oligomers. One electrode was configured to translate over anelectrode surface at a constant gap as illustrated in FIG. 63. Thetypical speed of translation was about 8.6 nm/s, which simulates DNAbeing pulled through, for example, a nanopore.

FIG. 64 shows exemplary current distributions for d(CCCCC) (FIG. 64A)and d(AAAAA) (FIG. 64B). The results of sequential reads for,homopolymers are summarized in FIGS. 65-68 (AAAAA—FIG. 65; CCCCC—FIG.66; d(^(m)C)₅—FIG. 67; and d(CCCCC) (FIG. 68)), plotting the current(nA) by time (s). The reads display consistent signals for thehomopolymers.

The results of sequential reads for heteropolymers are summarized inFIGS. 70-75 (ACACA—FIG. 70; CCACC—FIG. 71; C^(m)CC^(m)CC—FIG. 72;d(ACACA)—FIG. 73; d(C^(m)CC^(m)CC)—FIG. 74); and d(GTCGTCGTC)—FIG. 75),plotting the current (nA) by time (s). The reads display consistentsignals for the heteropolymers.

Thus, the above tests verify that imidazole-2-carboxyamide is aneffective reagent for analyzing oligomers.

Example 22 Synthesis of 4-carbamonylphenyldithiocarbamate

To a solution of 4-aminobenzamide (2 mmol, 272 mg) in DMF (1 mL), NaH(60% in mineral oil, 1.2 eq, 9.6 mg) and CS₂ (1.5 eq., 181.3 uL) weresuccessively added at 0° C. as shown below.

After 30 min at 0° C., the reaction mixture was warmed to roomtemperature and stirred for 84 h, and then warmed to 60° C. and stirredfor 4 h. After cooling to room temperature, the reaction mixture wasdiluted with ether (20 mL), filtered, washed with ether to give ayellowish powder 183 mg (yield 39%).

It will be appreciated that various of the above-disclosed and otherfeatures and functions, or alternatives thereof, may be desirablycombined into many other different systems or applications. Also,various presently unforeseen or unanticipated alternatives,modifications, variations or improvements therein may be subsequentlymade by those skilled in the art, and are also intended to beencompassed by the following claims.

1. A device for analyzing a polymer, said device comprising (a) a firstand second electrode that form a tunnel gap through which said polymercan pass; and (b) a first reagent attached to the first electrode and asecond reagent attached to the second electrode wherein the first andsecond reagents are each capable of forming a transient bond to a unitof the polymer, wherein a detectable signal is produced when thetransient bond forms.
 2. The device of claim 1, wherein the reagent anda width of the tunnel gap are configured to produce a specificdetectable signal for each monomer of the polymer when the first andsecond reagents form the transient bond to said unit of the polymer. 3.The device of claim 1, wherein the first and/or second electrodecomprises gold, carbon, platinum, graphene, or titanium nitride.
 4. Thedevice of claim 1, wherein the detectable signal produced is caused bythe binding of only one unit of the polymer in the tunnel gap.
 5. Thedevice of claim 1, wherein the tunnel gap has a width of about 1 toabout 4 nm.
 6. The device of claim 1, wherein the first and secondreagent are the same.
 7. The device of claim 1, wherein the transientbond is a hydrogen bond.
 8. The device of claim 1, wherein the first andsecond reagents comprise at least one hydrogen bond donor and at leastone hydrogen bond acceptor in an aqueous solution.
 9. The device ofclaim 1, wherein the first and second reagent are independently selectedfrom the group consisting of mercaptobenzoic acid,4-mercaptobenzcarbamide, imidazole-2-carboxide, and4-carbamonylphenyldithiocarbamate.
 10. The device of claim 1, whereinthe polymer is DNA or RNA and the unit is a nucleotide.
 11. The deviceof claim 1, wherein the unit is selected from the group consisting of A,C, T, G, and ^(m)C.
 12. The device of claim 1, wherein the polymer is aprotein and the unit is an amino acid.
 13. The device of claim 1,further comprising c) a first fluid reservoir and a second fluidreservoir separated by at least one nanopore through which the polymermay flow.
 14. The device of claim 13, further comprising a first drivingelectrode and a second driving electrode to apply an electrophoreticbias between the first fluid reservoir and the second fluid reservoir.15. The device of claim 13, further comprising a top contact electrodepositioned on top of the nanopore, wherein the nanopore is electricallyconductive.
 16. The device of claim 1 wherein the first and secondelectrodes are carbon nanotubes.
 17. A method of analyzing a polymer orpolymer unit, the method comprising: a) forming a transient bond betweena unit of the polymer and a first electrode functionalized with a firstreagent and between the unit of the polymer and a second electrodefunctionalized with a second reagent; b) detecting a detectable signalwhen the transient bonds form.
 18. The method of claim 17, wherein thefirst and second reagent are the same and are selected from the groupconsisting of selected from the group consisting of mercaptobenzoicacid, 4-mercaptobenzcarbamide, imidazole-2-carboxide, and4-carbamonylphenyldithiocarbamate.
 19. The method of claim 17, whereinthe polymer is DNA or RNA and the polymer unit is a nucleotide.
 20. Amethod of sequencing a polymer, the method comprising a) allowing a unitof the polymer to flow into a tunnel gap formed between a firstelectrode functionalized with a first reagent and a second electrodefunctionalized with a second reagent; b) forming a transient bondbetween a unit of the polymer and the first and second reagents; c)detecting a detectable signal when the transient bonds forms; and d)repeating steps a)-c) for each sequential unit of the polymer.
 21. Thedevice of claim 1, wherein the transient bond is pi-stacking betweenaromatic groups in the reader molecule and aromatic groups in theanalyte.
 22. A device for slowing the speed with which anelectrophoretically-driven polymer passes through a nanopore consistingof molecules that are chemically bonded to the inside of the pore andthat form transient bonds with units of the polymer as they pass throughthe pore.